Practical capillary electrophoresis

PREFACE TO THE FIRST EDITION

Capillary electrophoresis (CE) or high-performance CE (HPCE) is making the transition from a laboratory curiosity to a maturing microseparations technique. Now used in almost 1000 laboratories worldwide, CE is employed in an ever-widening scope of applications covering both large and small molecules.

The inspiration for this book arose from my popular American Chemical Society short course entitled, as is this text, "Practical Capillary Electrophoresis." During the first 18 months since its inception, nearly 500 students have enrolled in public and private sessions in the United States and Europe.

I have been amazed at the diversity of the scientific backgrounds of my students. Represented in these courses were molecular biologists, protein chemists, analytical chemists, organic chemists, and analytical biochemists from industrial, academic, and government laboratories. Interestingly enough, CE provides the mechanism for members of this multidisciplinary group to actually talk with each other, a rare event in most organizations.

But the diverse nature of the group provides teaching challenges as well. Most of the students are well versed in the art and science of liquid chromatography. However, CE is not chromatography (usually). It is electrophoresis, and it is governed by the art and science of electrophoresis. For those skilled in electrophoresis, CE offers additional separation opportunities that are not available in the slab-gel format. Furthermore, the intellectual process of methods development differs from that in either slab-gel electrophoresis or liquid chromatography.

The key to grasping the fundamentals of CE is to develop an understanding of how ions move about in fluid solution under the influence of an applied electric field. With this background, it becomes painless to wander through the elec-trophoretic domain and explain the subdeties and permutations frequently illustrated on the electropherograms. Accordingly, a logical approach to methods

XIV Preface

development evolves from this treatment. This is the goal of my course, and hopefully, I have translated this message into this text.

Since I work independently, without academic or industrial affiliations, the writing of this text would have been impossible without the help of my friends and colleagues. In particular, 1 am grateful to Professor Ira Krull and his graduate student, Jeff Mazzeo, from Northeastern University for reviewing the entire manuscript; Dr. Michael Albin from Applied Biosystems, Inc., for providing his company's computerized bibliography on HPCE; and the Perkin-Elmer Corporation including Ralph Conlon, Franco Spoldi, and librarian Debra Kaufman and her staff for invaluable assistance. I am also thankful to my associates throughout the scientific instrumentation industry for providing information, intellectual challenges, hints, electropherograms, comments, etc., many of which are included in this text. Last, I thank my students for helping me continuously reshape this material to provide clear and concise explanations of elec-trophoretic phenomena.

Finally, many of the figures in this text were produced by scanning the illustration in a journal article with subsequent graphic editing. While all efforts were made to preserve the integrity of the original data, subtle differences may appear in the figures produced in this book.

Robert Weinberger Chappaqua, NY

August 1992

PREFACE TO THE SECOND EDITION

It is hard to believe that seven years have passed since I wrote the first edition of this book. The time is ripe for a second edition. Not only has capillary electrophoresis matured, but my ability to articulate the field has improved as well.

I have reorganized this book to better reflect usage in the field. There are now ten chapters instead of twelve. The material on isotachophoresis has been combined with the section on stacking, and the special topics chapter has been eliminated. With the exception of the introduction and the chapter on basic concepts, all of the other material has been extensively reorganized and rewritten. Emphasis has been placed on commercially available apparatus and reagents, although gaps in the commercial offerings are discussed as well. Note that micellar electro-kinetic capillary chromatography (MECC) is considered as a variant of capillary zone electrophoresis (CZE) and is included in the chapter on secondary equilibrium. Cyclodextrins and chiral recognition are covered here as well.

Many thanks to Dr. Bruce McCord, Mr. Ira Lurie, and Professor Ira KruU for reviewing some of the chapters in this second edition. The author gratefully acknowledges the support of Hewlett-Packard and in particular Dr. David Heiger.

Much has been said about the ability of capillary electrophoresis (HPCE) to replace liquid chromatography (HPLC). Clearly it has not. As the first high-performance condensed phase technique, HPLC quickly replaced gas chromatography as the method of choice for separating polar molocules. As food for thought, imagine if capillary electrophoresis had a 25-year head start over HPLC. Then perhaps the chromatographers would be fighting the uphill battle of displacing HPCE. As noted in this text, HPCE is clearly superseding the slab gel, at least in the fields of DNA separations.

Robert Weinberger Chappaqua, NY

June 1,1999

MASTER SYMBOL LIST

A Corrected peak area corr r

A Raw peak area raw r

a Fraction ionized a Molar absorptivity a Separation factor h Detector optical pathlength C, c Concentration C Coefficient for resistance to mass transfer in the mobile phase C^ Coefficient for resistance to mass transfer in the stationary phase CLOD Concentration limit of detection CMC Critical micelle concentration %C Percentage of crosslinker in a gel D Capillary diameter D, D Diffusion coefficient

m

D^^ Solute diffusion in stagnant mobile phase DR Dynamic reserve d Particle diameter, chromatography AH Height differential between capillary inlet and outlet Ap^ Difference in mobility between two solutes AP Pressure drop 6 Debye radius ^ Zeta potential e Charge per unit area E Field strength E Acceptable increase in H E Detector efficiency 8 Dielectric constant 8 Molar absorptivity 8o Permittivity of vacuum

XVi Master Symbol List

/ Frictional force (Stoke's law) g Gravitational constant Y Field enhancement factor Y Obstructive factor for diffusion, Van Deemter equation H Height equivalent of a theoretical plate dH/dt Rate of heat production I Current If Fluorescence intensity I Excitation source intensity k Conductivity k' Capacity factor k' Capacity factor in MECC K, X Thermal conductivity K Equilibrium constant L Length of capillary L Length of capillary to detector I Length of the detector window Lf Length of capillary from detector to fraction collector L ^ Length of the unpacked portion of a CEC capillary L ^^^^^ Length of the packed portion of a CEC capillary L Total length of capillary l.^. Length of an injection plug X Tortuosity factor, Van Deemter equation m Mass M Actual mass MLOD Mass limit of detection N Number of segments in a polymer chain N Number of theoretical plates n Number of charges r\ Viscosity P Partition coefficient between water and micelle

wm

AP Pressure drop O Polymer concentration, size separations O Quantum yield O Overlap threshold Oj Fluorescence quantum yield O* Entanglement threshold, size separations p Density p Resistivity Q Quantity of injected material q Ionic net charge R Resistance R Peak ratio R Displacement ratio

Master Symbol List XVll

R Resolution s

r Ionic radius (Stokes' law) r Capillary radius S/N Signal to noise ratio a Peak variance a Peak variance due to capillary wall effects

cap ^ J

a^ Peak variance due to the detector det

a, „ Peak variance due to diffusion diff

a ^ Peak variance due to electrodispersion heat P^ak variance due to Joule heating

a Peak variance due to injection mj J

a^ Peak variance in units of length o Peak variance from all sources

tot

Time Absorption time to a stationary phase or wall Desorption time from a stationary phase or wall Lag time Migration time Migration time for a micellar aggregate Migration time for a neutral "unretained" solute Retention time

T Temperature TR Transfer ratio %)T Percentage of monomer and crosslinker in a gel

0/

|JL Ionic mobility |i^ Apparent (measured) mobility |Li^^ Electroosmotic mobility |Li^ Electrophoretic mobility V Partial molar volume of micelle V Voltage 1) Ionic velocity 1) Mean linear velocity D Electrophoretic velocity 1) Electroosmotic velocity

eo J

0) ^ Solute velocity in the unpacked portion of a CEC capillary \) ^^^^^ Solute velocity in the packed portion of a CEC capillary W Power W. Width of an injection plug W Spatial width of a sample zone W Temporal width of a sample zone X. Intital length of an injection plug X Zone length after stacking Z Number of valence electrons Z Charge

CHAPTER 1 Introduction

1.1 Electrophoresis 1.2 Microchromatographic Separation Methods 1.3 Capillary Electrophoresis 1.4 Capillary Electrochromatography 1.5 Micromachined Electrophoretic Devices 1.6 Historical Perspective 1.7 Generic HPCE Systems 1.8 Instrumentation 1.9 Sources of Information on HPCE 1.10 Capillary Electrophoresis: A Family of Techniques

References

1.1 ELECTROPHORESIS

Electrophoresis is a process for separating charged molecules based on their movement through a fluid under the influence of an applied electric field. If two solutes have differing electrophoretic mobilities, then separation v^U usually occur. The separation is performed in a medium such as a semisolid slab-gel. Gels provide physical support and mechanical stabiUty for the fluidic buffer system. In some modes of electrophoresis, the gel participates in the mechanism of separation by serving as a molecular sieve. Nongel media such as paper or cellulose acetate are alternative supports. These media are less inert than gels, as they contain charged surface groups that may interact with the sample or the run buffer.

A carrier electrolyte is also required for electrophoresis. Otherwise known as the background electrolyte (BGE), the carrier electrolyte, or simply the run buffer, this solution maintains the requisite pH and provides sufficient conductivity to allow the passage of current (ions), necessary for the separation. Frequently, additional materials are added to the BGE to adjust the resolution of the separation through the generation of secondary equilibria. Additives can also serve to maintain solubility and prevent the interaction of solutes or excip-ients with the gel matrix or, in the case of capillary electrophoresis, with the

^ Chapter 1 Introduction

capillary wall. The theory and practice of electrophoresis have been the subject of many textbooks and conference proceedings (1-9).

Apparatus for conducting electrophoresis, such as that illustrated in Figure 1.1, is remarkably simple and low cost. The gel medium, which is supported on glass plates, is inserted into a Plexiglass chamber. Two buffer reservoirs make contact at each end of the gel. Electrodes immersed in the buffers complete the electrical circuit between the gel and power supply. Many samples can be separated simultaneously, since it is possible to use a multilane gel. One or two lanes are frequently reserved for standard mixtures to calibrate the electropherogram. Calibration is usually based on molecular size or, in isoelectric focusing, pi.

Gels such as polyacrylamide or agarose serve several important functions:

1. they may contribute to the mechanism of separation; 2. they reduce the dispersive effects of diffusion and convection; and 3. they serve to physically stabilize the separation matrix.

The gel composition is adjusted to define specific pore sizes, each for a nominal range of molecular sizes. This forms the basis for separations of macro-molecules based on size. By proper calibration, extrapolation to molecular weight is straightforward.

Reduction of convection and diffusion is an important function of the gel matrix. The production of heat by the applied field induces convective movement of the electrolyte. This movement results in band broadening that reduces the efficiency of the separation. The viscous gel media inhibits fluid movement in the electric field. Such a material is termed anticonvective. Since the gel is of high viscosity, molecular diffusion is reduced as well, further enhancing the efficiency of the separation.

CATHODE

ANODE

BUFFER SOLUTION

GEL

BUFFER SOLUTION

FIGURE 1.1 Drawing of an apparatus for slab-gel electrophoresis.

1.2 Microchromatographic Separation Methods 3

Finally, the gel must be sufficiently viscous to provide physical support. Low viscosity solutions or gels would flow if the plate is not held level. Immersion in detection reagents would be impossible, since handling or contact with fluid solutions would destroy the matrix and separation. In the capillary format, the gel is unnecessary since the walls of the capillary provide the mechanical stability for the separation.

The basic procedure for performing gel electrophoresis is as follows:

1. prepare, pour and polymerize the gel; 2. apply the sample; 3. run the separation; 4. immerse the gel in a detection reagent; 5. destain the gel; 6. preserve the gel; and 7. photograph or scan the gel for a permanent record.^

These steps are extremely labor intensive. High performance capillary electrophoresis (HPCE) is the automated and instrumental version of slab-gel electrophoresis. In the DNA applications arena, the most important of which include DNA sequencing, human identification, and genetic analysis, HPCE is rapidly replacing the slab-gel as the separation method of choice.

The separation of some polymerase chain reaction (PCR) products is shown in Figure 1.2. A restriction digest, used as a sizing standard, appears in the outer lanes. The middle three lanes of the gel show a triplicate run of a 500-mer double-stranded DNA PCR reaction. Quantitation for such a separation is difficult and often imprecise, but such information can be obtained with the aid of a gel scanner. Recoveries of material from the gel are performed using procedures such as the Southern blot (10). Sufficient material is recoverable for sequencing or other bioassays.

Separations of the sizing standard and 500-mer PCR product by HPCE using a size selective polymer network are shown in Figure 1.3. Quantitation is readily performed using peak area comparison with the standard. However, fraction collection is difficult relative to the slab-gel, particularly for trace impurities, since only minuscule amounts of material are injected into the capillary.

1.2 MICROCHROMATOGRAPHIC SEPARATION METHODS

The evolution of chromatographic methods over the last 40 years has produced a systematic and rational trend toward miniaturization. This is particularly true

lOn-line detection is performed on an instrument such as an automated DNA sequencer. ^Automated gel scanners can be used in place of gel archiving or photography.

Chapter 1 Introduction

If

FIGURE 1.2 Slab-gel electrophoresis of a 500-mer double-stranded PCR reaction product in a 1.8% agarose ethidium bromide gel. Courtesy of Bio-Rad.

for gas chromatography, where the advantages of the open tubular capillary displaced the use of packed columns for most applications.

Chromatographic separations all function via differential partitioning of a solute between a stationary phase and a mobile phase. A packed column offers solutes "a multiplicity of flow paths, some short, the majority of average length, and some long (11)." Solute molecules select various paths through the chromatographic maze. The detected peak suggests this distribution and is broadened. In the open tubular capillary, the choices for solute transport are limited, so that the solute elutes as a narrow band.

In order for the open tubular capillary to function properly, its diameter must be quite small. Larger diameter capillaries present a problem, since solutes away from the walls do not sense the stationary phase in a timely fashion. However, a major problem with narrow inner diameter (i.d.) capillaries is loading capacity. Injection sizes must be kept small to avoid overloading the system. In gas chromatography (GC) this problem is overcome in part, since sensitive detectors such as the flame ionization detector (FID), electron capture detector (ECD), and mass spectrometer are easily interfaced.

Improved efficiency is one of several advantages obtained through miniaturization. The most important of those is improved mass limits of detection

1.2 Microchromatographic Separation Methods

i 500

IL 1746

303 ^^

10 15 TIME (min.)

20

FIGURE 1.3 Capillary gel electrophoresis of a 500-mer (top) double-stranded PCR reaction product and a low molecular weight sizing standard (bottom). Capillary: 50 cm x 50 [im i.d. Bio-Rad coated capillary; buffer: 100 mM tris-borate, pH 8.3, 2 mM EDTA with linear polymers; injection: electrokinetic, 8 kV, 8 sec; detection UV, 260 nm. Courtesy of Bio-Rad.

(MLOD). Since dilution of the solute is minimized in the miniaturized system, better MLODs are obtained than in large scale systems. This is particularly important when the available sample size is small, as sometimes happens in bio-molecule separations.

Miniaturization of GC has been exquisitely successful. These triumphs could not be directly transferred to liquid chromatography (LC) for several reasons. The most important is the lack of good detectors. Interface to the FID and ECD is not practical due to the incompatibility of the mobile phase with each detector. Pumping of the mobile phase at the low flow rates required by miniaturization is also more complex, particularly when gradient elution is required. Despite these problems, |I-LC systems are useful in sample-limited situations and for mass spectrometry where the reduced liquid flow rate is advantageous. Several books have been devoted to this important field (12-14).

6 Chapter 1 Introduction

Most of work with \i-LC employs 250 |im i.d. packed columns, and so the advantages enjoyed by open tubular GC are not realized in |Li-LC. The instrumental problems of injection and detection posed by open tubular LC have inhibited most people from using this technology

1.3 CAPILLARY ELECTROPHORESIS

The arrival of HPCE solved many experimental problems of gels. Use of gels is unnecessary since the capillary walls provide mechanical support for the carrier electrolyte.3 The daunting task of automation for the slab-gel format is solved with HPCE. Sample introduction (injection) is performed in a repeatable manner. Detection is on-line, and the instrumental output resembles a chro-matogram. The use of narrow diameter capillaries allows efficient heat dissipation. This permits the use of high voltage to drive the separation. Since the speed of electrophoresis is directly proportional to the field strength, separations by HPCE are faster than those in slab-gels. On the other hand, the relative speed of the slab-gel is enhanced, since multiple samples can be separated at once. HPCE is a serial technique; one sample is followed by another. This limitation has been overcome through the use of the capillary array for high throughput applications such as DNA sequencing (15,16) and serum protein analysis (17). Commercial instruments are now available for these applications.

HPCE represents a merging of technologies derived from traditional electrophoresis and high performance liquid chromatography (HPLC). Both HPCE and HPLC employ on-line detection. Developments in on-column micro-LC detection have directly transferred over to capillary electrophoresis. One of the modes of HPCE, micellar electrokinetic capillary chromatography (Chapter 4), can be considered a chromatographic technique. Electrically driven separations through packed columns (Chapter 7) have been reported from many laboratories. While there is much in common between chromatography and electrophoresis, the fundamentals of HPCE are based on electrophoresis, not chromatography.

Professor Richard Hartwick, formerly from the State University of New York at Binghamton, started many of his lectures on capillary electrophoresis with a discussion of transport processes in separations. While performing a separation, there are two major transport processes occurring:

Separative transport arises from the free energy differences experienced by molecules with their physicochemical environment. The separation mechanism may be based on phase equilibria such as adsorption, extraction, or ion exchange. Alternatively, kinetic processes such as electrophoresis or dialysis provide the mechanism for separation. Whatever the mechanism for separation, each individual solute must have unique transport properties for a separation to occur.

^Gels are occasionally used in HPCE for running size separations. Pumpable polymer networks are preferred, since they can be changed for each run.

1.3 Capillary Electrophoresis 7

Dispersive transport, or band broadening, is the sum of processes of the dispersing zones about their center of gravities. Examples of dispersion processes are diffusion, convection, and restricted mass transfer. Even under conditions of excellent separative transport, dispersive transport, unless properly controlled, can merge peaks together.

According to the late Professor Calvin Giddings as paraphrased by Hartwick, "separation is the art and science of maximizing separative transport relative to dispersive transport." In this regard, capillary electrophoresis is perhaps the finest example of optimizing both transport mechanisms to yield highly efficient separations.

Figures 1.4 and 1.5 illustrate this concept, using a series of barbiturate separations to compare HPCE and HPLC. The mode of electrophoresis used in Figure 1.5 is micellar electrokinetic capillary chromatography (MECC), an electrophoretic technique that resembles reversed-phase LC. In the LC separation amobarbital and pentabarbital coelute, but they are resolved by HPCE.

With some optimization work, amobarbital and pentabarbital can be separated by HPLC. But with HPCE, methods development often progresses rapidly because of the enormous peak capacity of the technique. Peak capacity simply describes the number of peaks can be separated per unit time. With a couple of hundred thousand theoretical plates,"^ many separations occur without extensive optimization efforts. In addition, peak symmetry is excellent using HPCE unless wall effects (Section 3.3) occur. With the absence of a stationary phase, many factors that contribute to peak broadening and tailing are minimized.

It would be misleading to state that all separations are superior by HPCE or that methods development will always be straightforward. It is realistic, however, based on the experiences of many separation scientists skilled in the art of both techniques, to predict that HPCE will provide the requisite speed and resolution in the shortest possible run time with the least amount of methods development, under most circumstances.

These same two figures illustrate an important limitation of HPCE, the concentration limit of detection (CLOD). In Figure 1.4, the LC separation requires a 1.25 |Lig/mL solution to give full scale peaks with 1-2% noise (the postcolumn reagent merely alkalized the mobile phase, permitting sensitive detection at 240 nm). The CLOD is approximately 30-fold better by HPLC. The MECC separation shown in Figure 1.5 required a solute concentration of 100 |ag/mL for a similar response, although the noise was lower (0.5%).^ On the other hand, the MLOD by capillary electrophoresis exceeds HPLC by a factor of 100. The ideal detector for HPCE will be mass sensitive and not depend on the narrow optical pathlength defined by the capillary itself. Descriptions, advantages, and limitations of many HPCE detectors can be found in Chapter 9.

^The theoretical plate (N) is a measure of the efficiency of a chromatographic of electrophoretic peak; N = 5.5'\(t^/Wiy, where t^ is the migration time and W is the peak width at half height. 5The CLOD can easily be improved through the use of stacking and/or extended pathlength flowcells.


W^ wW u TIME (MIN.) 11

FIGURE 1.4 Reversed-phase liquid chromatography of barbiturates. Column: Econosphere Cis, 25 cm X 4.6 mm i.d.; mobile phase: acetonitrile : water, 55/45 (v/v); injection size: 20 jxL; flow rate: 1.2 mL/min; postcolumn reagent: borate buffer, pH 10, 0.2 mL/min; detection: UV, 240 nm; solutes: (1) barbital, (2) butethel, (3) amobarbital and pentabarbital, (4) secobarbital; amount injected: 25 ng of each barbiturate from a 1.25 |Llg/mL solution.

The preceding comparison is significant since a |Li-separation technique is compared with conventional HPLC using a 4.6 mm i.d. column. Would it be better to compare HPCE with |i-LC? Perhaps so from an academic standpoint, but this would not reflect the current usage and thinking in the real world. Chemists are contemplating using HPCE to replace or augment conventional HPLC as well as |i-LC. Table 1.1 provides a comparison of slab-gel electrophoresis, |I-LC, HPLC, and HPCE. Two disadvantages of HPCE compared to conventional HPLC are sensitivity of detection and precision of analysis. These have prevented the most widespread use of HPCE. On the other hand, HPCE is replacing the slab-gel for most high-throughput DNA applications. In this case, the ease of automation, precision and ruggedness of HPCE supercede the slab-gel.

1.3 C apillary Electrophoresis

X TIME (MIN.) 10

FIGURE 1.5 Micellar electrokinetic capillary chromatography of barbiturates. Capillary: 50 cm (length to detector) X 50 |lm i.d.; buffer: 110 mM SDS, 50 mM borate, pH 9.5; injection: 1 sec vacuum (5 nL); detection: UV, 240 nm; solutes: (1) phenobarbital, (2) butethel, (3) barbital, (4) amo-barbital, (5) pentobarbital, (6) secobarbital; amount injected: 500 pg of each barbiturate from a 100 |lg/mL solution.

HPCE is a novel and alternative format for both liquid chromatography and electrophoresis. The unique properties of this technique include the use of:

1. capillary tubing in the range of 25-100 jim; 2. high electric field strength; 3. on-line detection in real time; 4. only nanoliters of sample; 5. limited quantities of mostly aqueous reagents; and 6. inexpensive capillaries relative to HPLC columns.

The molecular weight range of analytes separable by HPCE is enormous. A search of the literature reveals applications covering small ions, small molecules,


TABLE 1.1 Comparison of Slab-Gel Electrophoresis, p-LC, Conventional LC, and HPCE

Speed

Intrumentation cost

Sensitivity

CLOD

MLOD

Efficiency

Automation

Precision

Quantitation

Selectivity

Methods development

Reagent consumption

Preparative mode

Ruggedness

Separations

DNA Proteins Small molecules

Slab-Gel

slow

low

poor

poor

moderate

Htde

poor

difficult

moderate

slow

low

good

good

excellent excellent poor

p-LC

moderate

high

poor

good

moderate

yes

good

easy

moderate

moderate

low

fair

good

fair good excellent

HPLC

moderate

moderate

excellent

poor

moderate

yes

excellent

easy

moderate

moderate

high

excellent

excellent

fair good excellent

HPCE

fast

moderate

poor

excellent

high

yes

good

easy

high

rapid

minimal

poor

good

excellent excellent excellent

peptides, proteins, DNA, viruses, bacteria, blood cells, and colloidal particles. The molecular weight range of HPCE is easily from 3 for a lithium ion to 100,000,000 for a virus or particle.

1.4 CAPILLARY ELECTROCHROMATOGRAPHY

A hybrid of chromatography and electrophoresis, capillary electrochromatog-raphy (CEC) employs the electrically driven electroosmotic flow (EOF) to pump a mobile phase through a packed capillary. The use of the EOF to generate flow solves some of the instrumental problems of pumping at nL flow rates.

Capillary electrochromatography employs small diameter capillaries filled with a stationary phase. Reversed-phase packings are most often used, although an application with a cation-exchange material has been reported (18). An amazing efficiency 8 million plates per meter was reported in that paper, though the mechanism and reproducibility of the effect are still unclear.

1.6 Historical Perspective 11

Typically, 50 |im i.d. capillaries are used though larger diameter tubes can be employed at the expense of efficiency. Particle diameters of 3-5 |im porus material are most common, though it is possible to employ 1.5 |Lim pellicular packing. Since there is no pressure drop with an electrically pumped system, relatively long capillaries can be employed to generate hundred of thousands of theoretical plates. The reduction of eddy diffusion also contributes to the enhanced efficiency (19).

The mobile phase is pumped using the EOF generated by both the wall of the capillary and the chromatographic packing. Formulation of the mobile phase is similar to conventional reversed-phase chromatography, except that a dilute buffer—for example, 1-10 mM tris, borate, or phosphate—is added to ensure sufficient electrical conductivity The capillary is usually pressurized to a few atmospheres to suppress bubble formation.

The least mature of the electrically driven techniques, CEC capillaries and second generation instruments are now available. One promise for this technique is the ability to employ the vast existing chromatographic database to speed methods development.

1.5 MICROMACHINED ELECTROPHORETIC DEVICES

Employing technology used in the fabrication of integrated circuits, it is now possible to create an electrophoretic apparatus on a chip (20-28). Designed for dedicated applications such as clinical analysis, genetic analysis, or DNA sequencing, chips can be manufactured at low cost in commercial quantities. These devices can form the basis of an automated laboratory, where the disposable chip serves as the separations device.

A diagram of a simple micromachined HPCE chip is shown in Figure 1.6. The technological advantage of this device compared with a conventional capillary is its ability to perform extremely small injections (29). As a result, a shorter separation channel is required, again compared with the conventional capillary. Detection problems resulting from the small injection are solved through the use of laser-induced fluorescence (LIE). Micromachined electrophoretic devices are expected to have a huge impact in the DNA applications area.

1.6 HISTORICAL PERSPECTIVE

A century of development in electrophoresis and instrumentation has provided the foundation for HPCE. Reviews describing the history of electrophoresis were published by Vesterberg (30) and Compton and Brownlee (31). The highlights in the development of HPCE are given in Table 1.2.


Background Electrolyte

A B

oil o-

Sample

o-Separation Channel

Detector Window

FIGURE 1.6 Layout of the channels in a planar glass substrate. Channels are referred to by number and inlet points (reservoirs) as letters. Each channel is labeled with its content or its function. Overall dimensions are 14.8 cm x 3.9 cm x 1 cm thick. The location of one pair of platinum electrodes is shown; for clarity, the others are not. (A) BGE reservoir; (B) sample reservoir; (C) outlet reservoir. (1) BGE inlet; (2) sample inlet; (3) separation channel; (4) sample outlet. Injection is made where 4 crosses 3. Redrawn with permission from Anal. Chem., 64, 1926 (1992), copyright © Am. Chem. Soc.

A direct forerunner of modem CZE was developed by Hjerten in 1967 (32). To reduce the detrimental effects of convection caused by heat production, the 3 mm i.d. capillaries were rotated. While heat dissipation was unchanged, the rotating action caused mixing to occur within the capillary, smoothing out the convective gradients. In the 1970s, techniques using smaller i.d. capillaries were successfully developed (34). Superior heat dissipation permitted the use of higher field strength without the need for capillary rotation. In 1981, Jorgenson and Lukacs (35) solved the perplexing problems of injection and detection with 75 |Lim i.d. capillaries. Their advances clearly defined the start of the era of HPCE. Fluorescence detection was required at that time to record the electropherogram.

The 1980s proved ripe for invention. Adaptation of gel electrophoresis (36) and isoelectric focusing (38) to the capillary format was successful. In 1984, Terabe et al. (37) described a new form of electrophoresis called micellar elec-trokinetic capillary chromatography (MECC). Chromatographic separations of small molecules, whether charged or neutral, were obtained by employing the micelle as a "pseudo-stationary" phase.

Great advances in detection occurred during the 1980s to overcome, in part, the serious limitation of the short pathlength defined by narrow i.d. capillaries

1.6 H

istorical Perspective

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1 4 Chapter 1 Introduction

(57, 58). One of Jorgenson's first papers in the field employed fluorescence (59). Gassmann et al. (39) employed LIF, improving detectability to the attomole range. Olivares et al (42) interfaced CZE to the mass spectrometer via the elec-trospray. The use of on-line mass spectrometry is significant because of the difficulty of carrying out fraction collection. Wallingford and Ewing (43) developed electrochemical detection, sensitive enough to measure catecholamines in a single snail neuron. Kuhr and Yeung (44) employed indirect detection to measure solutes that neither absorbed nor fluoresced. More exotic detection techniques include electrochemical detection (43, 60) nuclear magnetic resonance (56), Raman (45), chemiluminescence (55) and radioactivity (61).

The problem of protein adherence to the capillary wall was addressed from several fronts. The use of treated capillaries was described by Hjerten (40) in 1985. Around the same time, Lauer and McManigill (41) employed alkaline buffers above the pi of the protein to effect solute repulsion from the anionic capillary wall. Based on these and related developments, wall effects have been substantially reduced.

The relative instability of crosslinked polyacrylamide gel-filled capillaries for protein and DNA separations was addressed by the first reports of polymer networks (47, 48). This led to the commercial introduction of kits for separations of proteins, oligonucleotides, and DNA. DNA sequencing can now be performed using various low viscosity polymer solutions (62, 63)

The first commercial instrument was introduced in 1988 by the late Bob Brownlee's company, Microphoretics. The following year, new instruments from Applied Biosystems, Beckman, and Bio-Rad were introduced. Later, Spectra-Physics, Isco, Europhor, Dionex, Waters Associates, Hewlett-Packard, and Uni-cam entered the fray. Modular systems from Lauer Labs, Groton Technologies, Jasco, and Europhor became available over the next few years.

In the mid 1990s, the slow development of the HPCE generic marketplace caused an industry shakeout as a number of instruments were withdrawn from the marketplace. In 1990, the first report employing a multiple capillary system was published (54). The mid to late 1990s provided the first application specific instruments for performing serum protein analysis (Beck-man) and DNA sequencing (Beckman, PE Biosystems, Molecular Dynamics). Instruments from the latter two companies are sold with 96 capillary arrays. These instruments are designed for high throughput DNA sequencing as required by the Human Genome Project. It is expected that human identification, another area that requires high throughput, will be implemented on these instruments.

In 1998, Covergant Bioscience Limited of Ontario, Canada, reported on a new dedicated instrument for capillary isoelectric focusing. The entire capillary is imaged using a charged coupled device camera. The advantage of whole capillary imaging is the elimination of the mobilization step. Electrochromatogra-phy has attracted intense interest in the late 1990s. The present state of the commercial offerings is given in Table 1.3.

1.6 H

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1.7 GENERIC HPCE SYSTEMS

While application specific DNA systems are becoming wildly successful, generic HPCE systems have not provided the returns expected by the scientific instrument manufacturers. The generic HPCE system is designed for the user to develop his or her own methods. In HPLC, this type of system forms the largest segment of this multibillion dollar market. There are numerous reasons, beyond the scientific, that this has not occurred with HPCE.

1. Liquid chromatography (HPLC) is the greatest analytical instrumentation success story in history. With a 23 year head start over HPCE, most problems have been worked out. Methods development is straightforward, chemists are trained, and troubleshooting is usually simple.

2. HPLC scales up for preparative work and scales down to the capillary format with relative ease. It is possible to have a single method for analytical, preparative, and commercial scale separations.

3. Many chromatographers consider HPCE to be the separations technique of last resort.

4. It is far more difficult for an instrument company sales force to sell HPCE. With quotas high and bonuses tied to performance, the salesperson goes where the money is. Setting up a demo instrument in a users lab is prone to failure, since the chemist is probably not trained in capillary electrophoresis. Postsales customer support is also quite high.

5. Capillary electrophoresis is electrophoresis, not chromatography. Chro-matographers must first master the principles of electrophoresis in order to effectively develop and troubleshoot methods. The training requirements are not trivial. Methods development can seem overwhelmingly complex to the new user.

6. Private industry is so downsized that scientists have no time to learn new techniques. Many purchased instruments sit idle because of initial failures of methods development. Instrument companies are downsized as well and have cut back on customer applications efforts. When faced with a problem chemists retreat to the familiar, and that is frequently HPLC.

7. Capillary electrophoresis is not as rugged as HPLC. Changes in the capillary surface chemistry lead to variable electroosmotic flow This in turn causes changes in the solute migration time.

8. The sensitivity of HPCE is lower than HPLC. This has become less of an issue as stacking techniques coupled with extended pathlength capillaries come into play. The training issues prevail, as many chemists are unaware of the variety of stacking techniques that exist.

9. There are few official methods of analysis employing HPCE. However, much is now in the pipeline. A number of pharmaceutical companies have submitted new drug applications to the Food and Drug Administration citing HPCE methodology.

1.8 Instrumentation 17

10. Since a single HPCE instrument can replace as many as ten liquid chroma tographs, the size of the market may become self-limiting.

The prospects for HPCE are not so bleak since once the learning curve is scaled. Successful methods development and routine implementation has been accomplished in many organizations.

1.8 INSTRUMENTATION

The instrumental configuration for HPCE is relatively simple. Before 1988, all work was done on simple homemade systems of a design similar to Jorgenson and Lukacs's original work (35, 59, 64).

A schematic of a homemade system is shown in Figure 1.7. The system consists of a high voltage power supply, buffer reservoirs, an HPLC ultraviolet detector, a capillary, and a Plexiglas cabinet. A safety interlock can be employed to prevent activation of high voltage when the cabinet is open. The capillary can be filled with buffer by a vacuum, generated using a syringe or handpump. Samples are injected either by siphoning (elevating the capillary for a defined time at a specified height) or by electrokinetic injection. While these simple systems provide good separations, precision may be poor due to the lack of temperature control and system automation.

Another common problem in homemade systems is excessive detector noise. The capillary is threaded through the detector and generally passes close to sensitive electronics, where the high electric field frequently causes electrical disturbances due

POWER SUPPLY

CAPILLARY

DETECTOR

CATHOLYTE ELECTRODES ^ ^ Q ^ ^ T E

(OUTLET) (INLET)

FIGURE 1.7 Basic schematic of an HPCE Instrument.


to inadequate grounding and shielding. This problem has been solved in commercial instrumentation.

The advantage of homemade systems is primarily in the area of detection. It is easy to interface HPCE to fluorescence detection and in particular laser-induced fluorescence. With the introduction of commercial modular systems, the advantages of homebuilt systems have all but disappeared excepting cost.

The arrival of commercial instruments has facilitated substantial growth in the field. An illustration of the now obsolete Applied Biosystems 270A is shown in Figure 1.8. This instrument provides the following basic features: a high voltage power supply that can provide up to 30 kV, an autosampler, electrodes, a separation capillary, an air-cooled capillary temperature controller, a UV detector, a capillary filling apparatus, and microprocessor control.

Newer instruments have random access where any vial can be designated as the inlet or outlet. Most of the newer instruments contain the capillary within a cartridge for efficient cooling with either air or fluids. Pressure is used rather than vacuum for filling the capillary; this is an advantage when using viscous polymers or interfacing to the mass spectrometer. Many instruments can also perform voltage programming and fraction collection, have alternative detectors such as fluorescence, photodiode array, or conductivity, and possess cooled

T Microprocessor

Reporting Integrator

Regulated Vacuum Reservoirs

Vent

Solenoid Valves

csfe

z Detector

1 Thermostated Compartment

Sample Buffer Reservoir

Sample Vials

Vacuum Buffer Reservoir Auto Sampler Carousel

FIGURE 1.8 Schematic of the Apphed Biosystems 270A. Courtesy of Apphed Biosystems.

1.9 Sources of Information on HPCE 1 9

autosamplers. Data systems that are specifically designed for HPCE are found on most units. Computers now provide for system control on all fully automated units.

1.9 SOURCES OF INFORMATION ON HPCE

Keeping up with the hterature in HPCE is no small task. Through 1998, about 7000 English language papers have appeared in the literature. The growth of the literature in the field is illustrated in Figure 1.9. Note the large increase that began in 1988, the year of commercial introduction of HPCE instrumentation.

The conference proceedings of the International Symposia on High Performance Capillary Electrophoresis that have appeared in the Journal of Chromatography, Vols. 480, 516,559,608, 652,680, 717, 744, 745, 781,817, and 853, contain an impressive concentration of state-of-the-art results. Other journals containing numerous papers on HPCE are Analytical Chemistry, Journal of Microcolumn Separations, Chromatographia, Journal of High Resolution Chromatography, Electrophoresis, Journal of Liquid Chromatography and Related Techniques, and Journal of Capillary Electrophoresis. Many dedicated issues from some of these journals covering HPCE, notably Electrophoresis and Journal of Liquid Chromatography, have been published as well.

For a comprehensive review of the literature, the biannual editions of Analytical Chemistry entitled "Fundamental Reviews" should be consulted. For general information on the theory of electromigration techniques, see (65) for an excellent review. Two recent editions of Electrophoresis, Vol. 18 (1997) No. 12-13 and Vol. 19 (1998) No. 16-17, contain outstanding

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FIGURE 1.9 The growth of the hterature of HPCE.


Table 1.4 Capillary Electrophoresis Books and Proceedings

Grossman, ED., Colbum, J.C., eds. Capillary Electrophoresis: Theory and Practice. 1992, Acadenic Press.

Vindevogel, J., Sandra, P Introduction to Micellar Electrokinetic Chromatography. 1992, Huthig.

Guzman, N., ed. Capillary Electrophoresis Technology. 1993, Marcel Dekker.

Weinberger, R. Practical Capillary Electrophoresis. 1993, Academic Press.

Camilleri, P., ed. Capillary Electrophoresis: Theory and Practice. 1993, CRC Press.

Foret, E, Krivankova, L., Bocek, P Capillary Zone Electrophoresis. 1993, VCH.

Jandik, P, Bonn, G. Capillary Electrophoresis of Small Molecules and Ions. 1993, VCH.

Baker, D. Capillary Electrophoresis. 1995, Wiley.

Righetti, P G., ed. Capillary Electrophoresis in Analytical Biotechnology. 1995, CRC.

Engelhardt, H., Beck, W, Schmitt, T. Capillary Electrophoresis: Methods and Potentials. 1996, Vieweg.

Cohen, A. S., Terabe, S., Deyl, Z., eds. Capillary Electrophoretic Separation of Drugs. 1996, Elsevier.

Altria, K.D., ed. Capillary Electrophoresis Guidebook: Principles, Operation, and Applications. 1996, Humana Press.

Jackim, E., ed. Capillary Electrophoresis Procedures Manual. 1996, Elsevier.

Lunte, S. M., Radzik, D. M., eds. Pharmaceutical and Biomedical Applications of Capillary Electrophoresis. 1996, Pergamon.

Coleman, D., ed. Directory of Capillary Electrophoresis: New Completely Revised Edition, 1996, Elsevier

Chankvetadze, B. Capillary Electrophoresis in Chiral Analysis. 1997, John Wiley 62: Sons.

Parvez, H., Caudy, P, Parvez, S., Roland-Gosselin, P, eds. Capillary Electrophoresis in Biotechnol

ogy and Environmental Analysis. 1997, VSP

Shintani, H., Polonsky J., ed. Handbook of Capillary Electrophoresis Applications. 1997, Blackie.

Weston, A., Brown, PR., HPLC and CE: Principle and Practice. 1997, Academic Press.

Heller, C , ed. Analysis of Nucleic Acids by Capillary Electro phoresis. 1997, Vieweg.

Khaledi, M.G., ed. High Performance Capillary Electrophoresis. Theory, Techniques, and Applications. 1998, Wiley

reviews of most aspects of HPCE. There have been numerous textbooks and conference proceedings in this field; a compilation is given in Table 1.4.

1.10 CAPILLARY ELECTROPHORESIS: A FAMILY OF TECHNIQUES

Capillary electrophoresis comprises a family of related techniques with differing mechanisms of separation. These techniques, which are covered in the following chapters of this book, are:

capillary zone electrophoresis (CZE) capillary isoelectric focusing (CIEF)

References 21

capillary gel electrophoresis (CGE)^ capillary isotachophoresis (CITP)-^ micellar electrokinetic capillary chromatography (MECC)^ capillary electroosmotic chromatography (CEC).

^CGE is now performed using replaceable polymer network reagents. 7CITP is considered here only for trace enrichment or sample stacking. ^MECC is the most significant application employing secondary equilibrium with CZE.

REFERENCES

1. Westheimer, R., Electrophoresis in Practice: A Guide to Methods and Applications ofDNA and Protein Separations, 2nd Ed. 1997, Wiley.

l.Mosher, R. A., Saville, D. A., Thormann, W, The Dynamics of Electrophoresis. 1992, VCH. 3.Rickwood, D., Hames, B. D., Gel Electrophoresis of Nucleic Acids: A Practical Approach, 2nd Ed.

1990, IRL Press. 4. Rickwood, D., Hames, B. D., Gel Electrophoresis of Proteins: A Practical Approach, 2nd Ed. 1990,

IRL Press. 5. Andrews, A. T., Electrophoresis: Theory, Techniques and Biochemical and Clinical Applications.

1981, Clarendon Press. 6. Righetti, P. G., Isoelectric Focusing: Theory, Methodology and Applications. Laboratory Techniques

in Biochemistry and Molecular Biology, ed. T. S. Work and R. H. Burdon. 1983, Elsevier Biomedical Press.

7. Chrambach, A., The Practice of Quantitative Gel Electrophoresis. 1985, VCH. S.Dunn, M. J., ed. Gel Electrophoresis of Proteins. 1986, Wright. 9.Jorgenson, J. W, Phillips, M., eds. New Directions in Electrophoretic Methods ACS Symposium

Series 335. 1987. American Chemical Society. 10.Southern, E. M.J. Mol. Biol, 1975; 98:503. 1 I.Jennings, W, Analytical Gas Chromatography. 1987, Academic Press, p.5. 12.Novotny, M., Ishii, D., eds. Microcolumn Separations. 1985, Elsevier. 13.1shii, D., ed. Introduction to Microscale High Performance Liquid Chromatography. 1988, VCH. 14. Yang, E J., ed. Microbore Column Chromatography: A Unified Approach to Chromatography. 1989,

Marcel Dekker. 15.Carrilho, E., Miller, A. W, Ruiz-Martinez, M. C, Kotler, L., Kesilman, J., Karger, B. L. Factors

to Be Considered for Robust High-Throughput Automated DNA Sequencing Using a Multiple-Capillary Array Instrument. Proc. SPIE-Int. Soc. Opt. Eng., 1997; 2985 (Ultrasensitive Biochemical Diagnostics II) :4.

16.Huang, X. C , Quesada, M. A., Mathies, R. A. Capillary Array Electrophoresis Using Laser-Excited Confocal Fluorescence Detection. Anal. Chem., 1992; 64:967.

17.Bienvenu, J., Graziani, M. S., Rpin, E A., Bernon, H., Blessum, C, Marchetti, C, Righetti, G., Somenzini, M., Verga, G., Aguzzi, E Multicenter Evaluation of the Paragon CZE 2000 Capillary Zone Electrophoresis for Serum Protein Electrophoresis and Monoclonal Component Typing. Clin. Chem., 1998; 44:599.

18. Smith, N. W, Evans, M. B. The Efficient Analysis of Neutral and Highly Polar Pharmaceutical Compounds Using Reversed-Phase and Ion-Exchange Electrochromatography. Chro-matographifl, 1995; 41:197.

19.Dittman, M. M., Wienand, K., Bek, E, Rozing, G. P. Theory and Practice of Capillary Electrochromatography LC-GC, 1995; 13:800.


20.Manz, A., Harrison, D. J., Verpoorte, E. M. J., Fettinger, J. C , Ludi, H., Widmer, H. M. Miniaturization of Chemical Analysis Systems—A Look into Next Century's Technology or Just a Fashionable Craze. Chimia, 1991; 45:103.

21.Manz, A., Harrison, D. J., Verpoorte, E. M. J., Fettinger, J. C, Paulus, A., Ludi, H., Widmer, H. M. Planar Chips Technology for Miniaturization and Integration of Separation Techniques into Monitoring Systems. Capillary Electrophoresis on a Chip. J. Chromatogr., 1992; 593:253.

22.Woolley, A. T, Mathies, R. A. Ultra-High-Speed DNA Sequencing Using Capillary Electrophoresis Chips. Anal. Chem., 1995; 67:3676.

23. Chiem, N. H., Harrison, D. J. Microchip Systems for Immunoassay: An Integrated Immunoreactor with Electrophoretic Separation for Serum Theophylline Determination. Clin. Chem., 1998; 44:591.

24. Colyer, C. L., Tang, T, Chiem, N., Harrison, D.J. Clinical Potential of Microchip Capillary Electrophoresis. Electrophoresis, 1997; 18:1733.

25.Effenhausen, C. S., Manz, A. Miniaturizing a Whole Analytical Laboratory Down to Chip Size. Am.Lah., 1994; 26:15.

26.Harrison, D. J., Manz, A., Fan, Z., Ludi, H., Widmar, H. M. Capillary Electrophoresis and Sample Injection Systems Integrated on a Planar Glass Chip. Anal. Chem., 1992; 64:1926.

27.Jacobson, S. C , Hergenroder, R., Koutny, L. B., Ramsey, M. J. High Speed Separations on a Microchip. Anal. Chem., 1994; 66:1114.

28.Jacobson, S. C , Hergenroder, R., Koutny, L. B., Ramsey, M.J. Open Channel Electrochro-matography on a Microchip. Anal. Chem., 1994; 66:2369.

29.Jacobson, S. C , Hergenroder, R., Koutny L. B., Warmack, R. J., Ramsey M. J. Effects of Injection Schemes and Column Geometry on the Performance of Microchip Electrophoresis Devices. Anal. Chem., 1994; 66:1107.

30.Vesterberg, O. History of Electrophoretic Methods. J. Chromatogr, 1989; 480:3. 31. Compton, S. W, Brownlee, R. G. Capillary Electrophoresis. BioTechniques, 1988; 6:432. 32.Hjerten, S. Free Zone Electrophoresis. Chromatogr Rev., 1967; 9:122. 33.Pretorius, V, Hopkins, B. J., Schieke, J. D. A New Concept of High-Speed Liquid Chromatogra

phy J. Chromatogr, 1974; 99:23. 34.Mikkers, F E. R, Everaerts, F M., Verheggen, T P. E. M. High Performance Zone Electrophore

sis. J. Chromatogr, 1979; 169:11. 35.Jorgenson, J. W, Lukacs, K. D. Zone Electrophoresis in Open Tubular Glass Capillaries. Anal.

Chem., 1981; 53:1298. 36.Hjerten, S. High-Performance Electrophoresis: The Electrophoretic Counterpart of High Per

formance Liquid Chromatography. J. Chromatogr, 1983; 270:1. 37.Terabe, S., Otsuka, K., Ichikawa, K., Tsuchiya, A., Ando, T. Electrokinetic Separations with

Micellar Solutions and Open-Tubular Capillaries. Anal. Chem., 1984; 56:111. 38.Hjerten, S., Zhu, M.-D. Adaptation of the Equipment for High-Performance Electrophoresis to

Isoelectric Focusing. J. Chromatogr, 1985; 346:265. 39.Gassmann, E., Kuo, J. E., Zare, R. N. Electrokinetic Separation of Chiral Compounds. Science,

1985; 230:813. 40.Hjerten, S. High-Performance Electrophoresis: Elimination of Electroendosmosis and Solute

Adsorption. J. Chromatogr, 1985; 347:191. 41.Lauer, H. H., McManigill, D. Capillary Zone Electrophoresis of Proteins in Untreated Fused Sil

ica Tubing. Anal. Chem., 1986; 58:166. 42.01ivares, J. A., Nguyen, N. T, Yonker, C. R., Smith, R. D. On-Line Mass Spectrometric Detec

tion for Capillary Zone Electrophoresis. Anal. Chem., 1987; 59:1230. 43. Wallingford, R. A., Ewing, A. G. Capillary Zone Electrophoresis with Electrochemical Detec

tion. Anal. Chem., 1987; 59:1762. 44.Kuhr, W G., Yeung, E. S. Indirect Fluorescence Detection of Native Amino Acids in Capillary

Zone Electrophoresis. Anal. Chem., 1988; 60:1832. 45. Chen, C. Y., Morris, M. D. Raman Spectroscopic Detection System for Capillary Zone Elec

trophoresis. Appl. Spectrosc, 1988; 42:515.

References 2 3

46. Guttman, A., Paulus, A., Cohen, A. S., Grinberg, N., Karger, B. L. Use of Complexing Agents for Selective Separation in High-Performance Capillary Electrophoresis: Chiral Resolution via Cyclodextrins Incorporated Within Polyacrylamide Gel Columns. J. Chromatogr., 1988; 448:41.

47.Hjerten, S., Valtcheva, L., Elenbring, K., Eaker, D. High-Performance Electrophoresis of Acidic and Basic Low-Molecular Weight Compounds and of Proteins in the Presence of Polymers and Neutral Surfactants. J. Liq. Chromatogr., 1989; 12:2471.

48.Zhu, M., Hansen, D. L., Burd, S., Gannon, F. Factors Affecting Free Zone Electrophoresis and Isoelectric Focusing in Capillary Electrophoresis. J. Chromatogr., 1989; 480:311.

49. Cohen, A. S., Najarian, D. R., Karger, B. L. Separation and Analysis of DNA Sequence Reaction Products by Capillary Gel Electrophoresis. J. Chromatogr, 1990; 516:49.

50.Drossman, H., Luckey J. A., Kostichka, A. J., D'Cunha, J., Smith, L. M. High-Speed Separations of DNA Sequencing Reactions by Capillary Electrophoresis. Anal. Chem., 1990; 62:900.

51.Luckey, J. A., Drossman, H., Kostichka, A. J., Mead, D. A., D'Cunha, J., Norris, T. B., Smith, L. M. High Speed DNA Sequencing by Capillary Electrophoresis. Nud. Acids Res., 1990; 18:4417.

52.Swerdlow, H., Gesteland, R. Capillary Gel Electrophoresis for Rapid, High Resolution DNA Sequencing. Nud. Adds Res., 1990; 18:1415.

53. Swerdlow, H., Wu, S., Harke, H., Dovichi, N. J. Capillary Gel Electrophoresis for DNA Sequencing: Laser-Induced Fluorescence Detection with the Sheath Flow Cuvette. J. Chromatogr, 1990; 516:61.

54.Zagursky, R. J., McCormick, R. M. DNA Sequencing Separations in Capillary Gels on a Modified Commercial DNA Sequencing Instrument. BioTechniques, 1990; 9:74.

55.Dadoo, R., Colon, L. A., Zare, R. N. Chemiluminescence Detection in Capillary Electrophoresis. HRC & CC, 1992; 15:133.

56. Wu, N., Peck, T. L., Webb, A. G., Magin, R. L., Sweedler, J. V Nanoliter Volume Sample Cells for ^H NMR: Application to Online Detection in Capillary Electrophoresis. J. Am. Chem. Soc, 1994; 116:7929.

57.Walbroehl, Y., Jorgenson, J. W On-Column UV Absorbance Detector for Open Tubular Capillary Zone Electrophoresis. J. Chromatogr, 1984; 315:135.

58. Green, J. S., Jorgenson, J. W Design of a Variable Wavelength UV Absorption Detector for On-Column Detection in Capillary Electrophoresis and Comparison of Its Performance to a Fixed Wavelength UV Absorption Detector. J. Liq. Chromatogr, 1989; 12:2527.

59.Jorgenson, J. W, Lukacs, K. D. Free-Zone Electrophoresis in Glass Capillaries. Clin. Chem., 1981; 27:1551.

60. Wallingford, R. A., Ewing, A. G. Capillary Zone Electrophoresis with Electrochemical Detection in 12.7|Lim Diameter Columns. Anal. Chem., 1988; 60:1972.

61.Pentoney, S. L., Zare, R. N., Quint, J. F. On-Line Radioisotope Detection for Capillary Electrophoresis. Anal. Chem., 1989; 61:1642.

62.Salas-Solano, O., Carrilho, E., Kolter, L., Miller, A. W, Goetzinger, W, Sosic, Z., Karger, B. L. Routine DNA Sequencing of 1000 Bases in Less than One Hour by Capillary Electrophoresis with Replaceable Linear Polyacrylamide Solutions. Anal. Chem., 1998; 70:3996.

63. Kim, Y., Yeung, E. S. Separation of DNA Sequencing Fragments up to 1000 Bases by Using PolyCethylene Oxide)-Filled Capillaries. J. Chromatogr, A, 1997; 781:315.

64.Jorgenson, J. W, Lukacs, K. D. Zone Electrophoresis in Open-Tubular Glass Capillaries: Preliminary Data on Performance. HRC & CC, 1981; 4:230.

65.Kleparnik, K., Bocek, P Theoretical Background for Clinical and Biomedical Applications of Electromigration Techniques. J. Chromatogr, 1991; 569:3.

CHAPTER 2 Capillary Zone Electrophoresis Basic Concepts

2.1 Electrical Conduction in Fluid Solution 2.2 The Language of Electrophoresis 2.3 Electroendoosmosis 2.4 Efficiency 2.5 Resolution 2.6 Joule Heating 2.7 Optimizing the Voltage and Temperature 2.8 Capillary Diameter and Buffer Ionic Strength 2.9 Optimizing the Capillary Length 2.10 Buffers 2.11 Temperature Effects 2.12 Buffer Additives 2.13 Capillaries 2.14 Sources of Band Broadening

References

2.1 ELECTRICAL CONDUCTION IN FLUID SOLUTION

Several simple concepts are important for understanding the physical processes that occur upon passage of an electrical current through an ionic solution. These processes are far more complex than the passage of current through a metal. In metals, uniform and v^eightless electrons carry all the current. In fluid

See any basic text on physical chemistry for a thorough description of electrical conduction in fluid solution.

25

26 Chapter 2 Capillary Zone Electrophoresis

solution, the current is carried by cations and anions. The molecular weight of these charge bearing ions ranges from a simple proton to tens of thousands for large complex ions such as proteins and polynucleotides.

Conduction in fluid solution is still described by Ohm's law,

E = IR, (2.1)

where E is the voltage or applied field, I is the current that passes through the solution, and R is the resistance of the fluid medium.

The reciprocal of resistance is conductivity. Kohlrausch found that the conductivity of a solution resulted from the independent migration of ions. As illustrated in Figure 2.1, when a current passes through an ionic solution, anions migrate toward the anode (positive electrode) while cations migrate toward the cathode (negative electrode) in equal quantities. Despite the passage of current, electroneutrality of the solution is always maintained because of electrolysis at each electrode.

This is important because electrolysis produces protons at the anode and hydroxide at the cathode (Figure 2.2). The resultant pH changes are due to the process known as buffer depletion (1-3). Since pH is the single most important experimental parameter in capillary electrophoresis, this effect must be minimized by

1. Using the appropriate buffers 2. Having sufficiently large buffer reservoirs 3. Replacing buffers frequently

The introduction of a sample into the capillary changes the situation dramatically (Figure 2.3). The Ohm's law equation changes as well to that for a series circuit:

E = IR^ + IRj . (2.2)

This process and the equation have important implications in HPCE. When low-conductivity samples (relative to the BGE) are injected, a process known as stacking (Section 8.6) occurs. This permits the use of large-volume injections

CATHODE ANODE

FIGURE 2.1 The independent migration of ions.

2.1 Electrical Conduction in Fluid Solution 27

POWER SUPPLY

CAPILLARY

DETECTOR OH GENERATED

AT CATHODE H GENERATED

AT ANODE

CATHOLYTE (OUTLET)

ELECTRODES

FIGURE 2.2 Buffer depletion.

ANOLYTE (INLET)

to be employed without excessive band broadening because zone compression occurs. On the other hand, if high-conductivity samples are injected relative to the BGE, antistacking or zone broadening will occur.

The conductivity of a solution is determined by two factors:

1. The concentration of the ionic species. 2. The speed of movement or mobility of the ionic species in an electric

field. In other words, highly mobile species are also highly conductive, and vice versa.

IR IR,

©,

FIGURE 2.3 Impact of the sample injection on the IR drops in a capillary.

2 8 Chapter 2 Capillary Zone Electrophoresis

The mobility of ions in fluid solution is governed by their charge to size ratio. The size of the molecule is based on the molecular weight, the three-dimensional structure, and the degree of solvation (usually hydration). Data given in Table 2.1 (4) for alkali metals illustrate several of these important points:

1. The orders for the mobilities of the metal ions are the reverse of what is expected based on the metal or crystal radii data. These smaller ions are more hydra ted than their larger counterparts.

2. The current generated by 100 mM solutions of various acetate salts is proportional to the ionic mobility of the cation. This feature becomes important when selecting the appropriate counterion for preparing buffer solutions.

The forces governing this behavior are expressed by Stoke's law,

/ = 67rrirv , (2.3)

where 7] = viscosity, r = ionic radius, and v = ionic velocity. The competing forces of mobility (velocity) and viscosity are illustrated in Figure 2.4 for an ion of radius r. Ionic size modifies mobility because of a solute's exposure to fric-tional drag as it migrates through the supporting electrolyte. The frictional drag is directly proportional to viscosity, size, and electrophoretic velocity. An expression for mobility that contains these terms is

M—)=-5^^^^^ = ^ - , (2.4) Vs E(V/cm) 6nrir

where q = the net charge and E = the electric field strength. Thus, mobility is considered a charge-to-size ratio. Since the units for velocity are centimeters per second and the field strength is expressed as volts per centimeter, the units of mobility are cm^A^s.

2.2 THE LANGUAGE OF ELECTROPHORESIS

There are several distinguishing differences between the terminology of chromatography and that of capillary electrophoresis. For example, a fundamental parameter in chromatography is the retention time. In electrophoresis nothing should ever be retained (except for CEC), so a more descriptive term is migration time: the time it takes a solute to travel from the beginning of the capillary to the detector window.

2.2 T

he L

angu

age of Electrop

horesis

29

< I—I

2 o

1

T1

<SJ

t 

»-l

3

0 u

fi o

2 »-

OS

,

^

B

X

0

TJ

!U

P^nd

OH

< ;>> o

<

"rn

3 T

l rr!

X ^

II U P

i

Sf2

hs. in

^ in

^

ON

O

rH

ON

ON

O

rn

00

d —

1 .-I

(N

s u

u =^

0 ^

r^ O

N

5 o ^j

<-

.


MOBILITY

FRICTIONAL FORCES

FIGURE 2.4 The competing forces of electrophoretic mobility and viscous drag.

The use of a detection window in HPCE (on-capillary detection) as opposed to postcolumn detection must also be considered. In HPLC, the length of the chromatographic column must be included in all methods. Figure 2.5 is a drawing of a capillary. Both the total length of the capillary (L or L) and the length to the detector (L^ or I) must be described. The segment of capillary that occurs after the detector window is necessary to make electrical contact with the outlet or detector-side electrolyte reservoir. Ideally, L^ - L^ should be as short as practical. Otherwise, some system voltage (V) is wasted on maintaining field strength (E) over part of the capillary that lies beyond the detector window and hence does not participate in the separation.

Lt

DETECTION WINDOW FIGURE 2.5 Illustration of a capillary defining the total length (Lj) and the length to the detector (Ld).

"ep

Mep E

.jjtrn

2.3 Electroendoosmosis 31

Expressions for some other fundamental terms are given in the following equations:

(2.5)

(2.6)

The preceding include the electrophoretic mobility (^ep^ cm^A^s), the elec-trophoretic velocity (Vgp, cm/s), and the field strength (E, V/cm). These equations define some fundamental features of HPCE:

1. Velocities are measured experimentally (Eq. 2.5). They are determined by dividing the length of capillary, from the injection side to the detector window (Ld), by the migration time t^.

2. MobiUties are calculated by dividing the electrophoretic velocity v p by the field strength (Eq. 2.6). The field strength is simply the voltage divided by the total capillary length (L ). The field strength is the important parameter governing electrophoretic migration. Field strength is changed when either the voltage or the capillary length is altered. Mobility is the fundamental parameter of capillary electrophoresis. This term is independent of voltage and capillary length. Equations (2.6) and (2.7) define only the relative mobility. To calculate the true mobility, a correction for a phenomenon known as electroendoosmotic flow (Section 2.3) must first be made.

2.3 ELECTROENDOOSMOSIS

A. THE CAPILLARY SURFACE

One of the fundamental processes that accompanies electrophoresis is electroos-mosis. One of the "pumping" mechanisms of HPCE, electroosmosis occurs because of the surface charge, known as the zeta potential, on the wall of the capillary. Fused silica is the most common material used to produce capillaries for HPCE. Technology developed for manufacturing capillary columns for GC readily transferred to HPCE. Fused silica is a highly crosslinked polymer of silicon dioxide with tremendous tensile strength (5), although it is quite brittle. With its polyimide coating, fused silica is quite durable, although some polyimide must


be removed to create a ultraviolet (UV) transparent optical window for detection. Other materials such as Teflon and quartz have been used (6), but performance and cost are less favorable.

Before use, capillaries are usually conditioned with 1 N sodium hydroxide. The base ionizes free silanol groups and may cleave some silica epoxide linkages as well. An anionic charge on the capillary surface results in the formation of an electrical double layer. The resulting ionic distribution is shown in Figure 2.6 (7). Anions are repelled from the negatively charged wall region, whereas cations are attracted as counterions. Ions closest to the wall are tightly bound and immobile, even under the influence of an electric field. Further from the wall is a compact and mobile region with substantial cationic character. At a greater distance from the wall, the solution becomes electrically neutral as the zeta potential of the wall is no longer sensed. Expressions describing this phenomenon were derived by Gouy and Chapman in 1910 and 1913, respectively This diffuse outer region is known as the Gouy-Chapman layer. The rigid inner layer is called the Stem layer.

When a voltage is applied, the mobile positive charges migrate in the direction of the cathode or negative electrode. Since ions are solvated by water, the fluid in the buffer is mobilized as well and dragged along by the migrating charge. Although the double layer is perhaps 100 A thick, the electroendoos-motic flow (EOF) is transmitted throughout the diameter of the capillary, presumably through hydrogen bonding of water molecules or van der Waals interactions between buffer constituents.

The electroosmotic flow as defined by Smoluchowski in 1903 is given by

Veo = ^ E , (2.8)

where £ is the dielectric constant, 77 is the viscosity of the buffer, and f is the zeta potential of the liquid-solid interface. The equation is only valid for capillaries sufficiently large that the double layers on opposite walls do not overlap each other (8). Practical use of this equation is not forthcoming, as the zeta potential is rarely measured and data for the dielectric constants of mixtures are not readily available. Like electrophoretic mobility, the EOF is inversely proportional to the viscosity of the BGE.

B. MEASURING THE ELECTROOSMOTIC FLOW

Since the migration time of a solute is influenced by the EOF, calculation of the actual mobility requires measurement of the EOF:

Here jH^^ = the actual mobility, jU pp = apparent (observed) mobility, and jLl^o = electroosmotic mobility. The use of mobility as the "migration parameter"

inttrfaisa

c . ^ g — : SI

®, ' "^ lA J Z*^^ /'""x x*i*'

I • X -

adsorbed compact diffuse layer layer layer

B

« 4N#

0 fit i

interf8€0

1 i s

1 1 u r L

<*•

1 1

^ l ^ \

compact. dlffuM***" layer 1 ''yr

' ^ •

HMliiiiiiliilnpi

dislanai from lh# eolynin wail FIGURE 2.6 Representation of the electrical double layer versus distance from the capillary wall. Reprinted with permission from J. Chromatogr., 559, 69 (1991), copyright © 1991 Elsevier Science Publishers.

will frequently yield greater precision compared to the use of migration time, since the impact of the EOF is factored out of the calculation (Section 10.6).

Routine measurement of the EOF is also necessary to ensure the integrity of the separation. If the EOF is not reproducible, it is likely that the capillary wall is being affected by some component in the sample or an experimental parameter is not being properly controlled (see Section 2.3F).


The simplest method for measuring the EOF is to inject a dilute solution containing a neutral solute and measure the time it takes to transit the detector (9-11). Since the capillary length is known, the velocity in centimeters per second is easily calculated. Dividing that value by the field strength yields the electroosmotic mobility in units of cmWs. Neutral solutes such as methanol, acetone, benzyl alcohol, and mesityl oxide are frequently employed. When MECC is the mode of separation (Chapter 4), a further requirement that the marker solute not partition into the micelle is imposed.

When the EOF is slow, the migration time can be quite long. To reduce the experimental time, it is useful to use the short end of the capillary to make the measurement. The short end is the section of capillary normally found between the detector window and capillary outlet. The injection can be made at the outlet side and the system operated using reversed polarity Now the EOF is measured using a short capillary length of 6-10 cm depending on the brand of instrument. As will be shown later, the short end of the capillary can be very useful when performing screening runs during methods development.

When the EOF is very slow, as in the case with certain coated capillaries, special techniques must be employed (12). It is seldom necessary to measure very weak EOF, since it does not notably affect mobility or experimental precision.

C. EFFECT OF BUFFER P H

The impact of pH on the EOF and the mobility is illustrated in Figure 2.7. At high pH the silanol groups are fully ionized, generating a strong zeta potential and dense electrical double layer. As a result, the EOF increases as the buffer pH is elevated (9, 13). A robust flow, typically around 2 mm/s at pH 9 in 20 mM borate buffer at 30 kV, 30°C is realized. For a 50 |Lim capillary, this translates to 235 nL/min. Since the total volume of a 50 cm x 50 jim i.d. capillary is only 980 nL, a neutral compound would reach the detector in 4.2 min. At pH 3, the EOF is much lower, about 30 nL/min.

The EOF must be controlled or even suppressed to run certain modes of HPCE. On the other hand, the EOF makes possible the simultaneous separation of cations, anions, and neutral species in a single run. For example, a zwitterion like a peptide will be negatively charged at a pH above its pi. The solute will elec-tromigrate toward the positive electrode. However, the EOF is sufficiently strong that the solute's net migration is toward the negative electrode (Figure 2.7, top). At low pH, the zwitterion has a positive charge and will migrate as well toward the negative electrode (Figure 2.7, bottom). In untreated fused-silica capillaries, most solutes migrate toward the negative electrode unless buffer additives or capillary treatments are used to reduce or reverse the EOF (Section 3.3).

The EOF is exquisitely sensitive to pH (9,14,15). Hysteresis effects have been reported (15) wherein the direction of approach to a particular pH value produces a different pH (Figure 2.8). When approaching from the acid side, the measured

2.3 Electroendoosmosis

HIGH pH

35

++++•'•+•'•+++++++++++++++++++++++++"'•+•*'+''"

M,0 / * ep 4 . 4 . ^ 4 . 4 , 4 » 4.4> 4. 4 . 4 . 4 . 4 . 4 . 4 . 4 . 4 * 4 .

+ + + + + 4* + 4- + + +^+ + -f + -I" + -f +

FIGURE 2.7 Behavior of electroendoosmotic flow and electrophoretic migration of a zwitterion (pi = 7) at high and low pH.

EOF is always lower, and vice versa. This means there is a kinetic parameter with regard to the estabUshment of a stable charge on the capillary wall. Longer equilibration times would reduce hysteresis at the expense of increased total run time.

Since the EOF will affect migration time precision, it is important to design experiments with these features in mind. The problems with EOF reproducibility are often most severe in the pH range 4-6 (15).

D. EFFECT OF BUFFER CONCENTRATION

The expression for the zeta potential is (16)

c = 47r5e

(2.10)

where £ = the buffer's dielectric constant, e = total excess charge in solution per unit area, and 5 is the double-layer thickness or Debye ionic radius. The Debye radius is 5 = (3 x 10'')(Z)(Ci/^), where Z = number of valence electrons and C = the buffer concentration.

As the ionic strength increases, the zeta potential and, similarly, the EOF decreases in proportion to the square root of the buffer concentration. This was


10

9 +

8 +

E o % 6 X u. o 5

4 +

3 +

H 1 1 1 1—I 1 1 1 1 2 3 4 5 6 7 8 9 10 11

PH

FIGURE 2.8 Effect of experimental design on the EOF. Key: • , high pH titrated to low pH; • , low pH titrated to high pH. Data from reference (15).

confirmed experimentally (17) for a series of buffers where the EOF was found hnear to the natural logarithm^ of the buffer concentration. It was reported that equivalent EOF is found for different buffer types as long as the ionic strength is kept constant (17).

The effect of buffer concentration and field strength is shown in Figure 2.9 (18). The electroosmotic mobility is plotted against field strength for phosphate buffer at three different concentrations using a 50-|Lim-i.d. capillary. As expected, the higher buffer concentrations showed lower EOF at all field strengths. Since

^The linear relationship of EOF with the buffer concentration is a square root relationship as indicated by Eq. (2.10).


71

u. ^^ u z o S

liini

UJ

50 100 150 200 E (V/cm)

250

FIGURE 2.9 Effect of buffer concentration and field strength (E, V/cm) on the electroosmotic flow in a 50-|xm-i.d. capillary. Buffer: phosphate at a concentration of (a) 10 mM; (b) 20 mM; (c) 50 mM. Redrawn with permission from J. Chromatogr., 516, 223 (1990), copyright © 1990 Elsevier Science Publishers.

mobility was plotted, all three lines should be flat. Slight positive slopes were reported for all three concentrations, presumably due to heating effects (Section 2.6). The same data produced using a lOO-jiim-i.d. capillary will be examined in that section.

E. EFFECT OF ORGANIC SOLVENTS

Organic solvents can modify the EOF because of their impact on buffer viscosity (17) and zeta potential (19). Linear alcohols such as methanol, ethanol, or isopropanol usually decrease the EOF because they increase the viscosity of the electrolyte. Acetonitrile either does not affect or may slightly increase the EOF (20). Organic solvents are often employed in HPCE to help solubilize the sample. Selectivity can be affected as well in both CZE (20) and MECC (21). Because of the sensitivity of organic solvent concentration on selectivity, evaporation must


be carefully controlled. In this regard, wholly aqueous separations are often advantageous.

F. CONTROLLING THE ELECTROOSMOTIC FLOW

The EOF is a double-edged sword. It allows the separation of cations, anions, and neutral solutes in a single run. It is also the single most important contributor to migration time variability on a run-to-run, day-to-day, and capillary-to-capillary basis. The EOF is affected by many parameters, including

Buffer pH

Buffer concentration

Temperature

Viscosity

Capillary surface

Field strength

Organic modifiers

Cellulose polymers

Surfactants

In this list, the only factor not under direct experimental control is the capillary surface. This single factor is often implicated as the cause for migration time variation in HPCE. It is important to ensure that the capillary surface is properly reconditioned after each run to maintain a reproducible surface. Coated capillaries that suppress the EOF are useful here, as long as the coating is stable. Some new reagents^ that form a dynamic surface coating show great promise for stabilizing the capillary surface (Section 3.3).

For some modes of HPCE, it is advantageous to suppress the FOE Capillary isoelectric focusing (CIEF) and capillary isotachophoresis (CITP) separations are usually performed under conditions of very low or carefully controlled EOF Additives such as 0.5% hydroxypropylmethyl cellulose are effective in suppressing the EOF, particularly in conjunction with a coated capillary (22). Cationic surfactants such as cetyltrimethylammonium bromide can actually reverse the direction of electroosmotic flow (14). This can be employed to prevent proteins from sticking to the capillary wall (23, 24). While complete suppression of the EOF is unnecessary for most applications, control is critical to obtain reproducible migration times and resolution.

^CElixir, Scientific Resources, Inc., Eatontown, NJ.

2.4 Efficiency 3 9

2.4 EFFICIENCY

The high efficiency of HPCE is a consequence of several unrelated factors:

1. A stationary phase is not required for HPCE. The primary cause of band broadening in LC is resistance to mass transfer between the stationary and mobile phases. This mass transfer problem is illustrated in Figure 2.10. When a solute is in the mobile phase, its linear velocity is determined by the linear velocity of the mobile phase. When attached to the stationary phase, the linear velocity becomes zero. The solute is not of a single velocity as it moves down the chromatographic tube. Whenever differing velocities occur during a separation, band broadening will occur. Minimizing the particle size of the packing improves but does not eliminate this problem. Thus, the parameter that results in separation also causes band broadening. The greater the retention, the greater the problem—as evidenced by broadened peaks as retention time increases. For most modes of HPCE (except CEC), this dispersion mechanism does not operate. Similarly, other HPLC dispersion mechanisms such as eddy diffusion and stagnant mobile phase are unimportant in HPCE.

2. In pressure-driven systems such as LC, the frictional forces of the mobile phase interacting at the walls of the tubing result in radial velocity gradients throughout the tube. As a result, the fluid velocity is greatest at the middle of the tube and approaches zero near the walls (Figure 2.11). This is known as laminar or parabolic flow. These frictional forces, together with the chromatographic packing, result in a substantial pressure drop across the column.

In electrically driven systems, the EOF is generated uniformly down the entire length of the capillary. There is no pressure drop in HPCE, and the radial flow profile is uniform across the capillary except very close to the walls, where the flow rate approaches zero (Figure 2.11).

FIGURE 2.10 The mass transport problem in HPLC.


Jorgenson and Lukacs derived the efficiency of the electrophoretic system from basic principles (25-27) using the assumption that diffusion is the only source of band broadening. Other sources of dispersion—including Joule heating (Section 2.6), capillary wall binding (Section 3.5), injection (Section 9.1), detection (Section 9.5), and electromigration dispersion (Section 2.13)—lead to fewer theoretical plates than the simple theory predicts.

The migration velocity for a solute is

V = luE = HV

(2.11)

where // = the mobility, E = field strength, V = voltage, and L = capillary length. The time t for a solute to migrate the length L of the capillary is

V fiV (2.12)

ELECTROOSMOTIC FLOW

HYDRODYNAMIC FLOW

FIGURE 2.11 Capillary flow profiles resulting from electroosmotic and hydrodynamic flow.

2.5 Resolution 41

Diffusion in liquids that leads to broadening of an initially sharp band is described by the Einstein equation

(yl = 2Dt = =^^, (2.13)

where D = the diffusion coefficient of the individual solute. The number of theoretical plates N is given by

N = — . (2.14)

Substituting Eq. (2.11) into Eq. (2.12) gives an expression for the number of theoretical plates:

N = ^ . (2.15) 2D

Some important generalizations can be made from this expression:

1. The use of high voltage gives the greatest number of theoretical plates, since the separation proceeds rapidly, minimizing the effect of diffusion. This holds true up to the point where heat dissipation is inadequate (Section 2.6).

2. Highly mobile solutes produce high plate counts, because their rapid velocity through the capillary minimizes the time for diffusion.

3. Solutes with low diffusion coefficients give high efficiency due to slow diffusional band broadening.

Points 2 and 3 appear contradictory. This is clarified by Figure 2.12 and supplemented with some calculations in Table 2.2. Because of the indirect but inverse relationship between mobility and diffusion, high-efficiency separations occur across a wide range of molecular weights.

HPCE can yield high-efficiency separations for both large and small molecules. The greatest number of theoretical plates is found in capillary gel electrophoresis (CGE). The use of an anticonvective gel matrix furthers the advantages of HPCE. The combination of HPCE in the gel or polymer network format (Chapter 6) can yield millions of theoretical plates.

2.5 RESOLUTION

While high efficiency is important, resolution is the key for all forms of separation. In a high-efficiency system, inadequate resolution may result in a single very sharp peak.


DIFFUSION

RAPID

SLOW

MOBILITY

HIGH

LOW

SMALL MOLECULES

LARGE MOLECULES

FIGURE 2.12 Diffusion and mobihty of small and large molecules.

The resolution (R) between two solutes is defined as

R. = 1 AAI^PVN

4 Mep + Meo

(2.16)

where A^ is the difference in mobility between two solutes, /i^^ is the average mobility of the two solutes, and N is the number of theoretical plates. Substituting the plate count equation (Eq. (2.15) and V = EL) yields (25)

R, = O.UlAfl^ EL

(/ ep + I^J^n (2.17)

This expression suggests that increasing the voltage is not very effective in improving resolution, since that parameter falls inside of the square root of the resolution equation. A doubling of voltage results in only a 41% improvement in resolution. The production of heat quickly limits this

Table 2.2 Calculated Theoretical Plates for a Small and Large Molecule

Solute

Horse heart myoglobin

Quinine sulfate

MW

13,900

747

Mobility (10-^ cmW-s)

0.65

4

Diffusion Coefficient

(10-^cmVs)

1

7

N

975,000

857,000

2.6 Joule Heating 43

approach. Another means of improving resolution as predicted by Eq. (2.17) is to adjust the EOF. Akhough this also falls under the square root sign of the resolution equation, this technique can be quite effective. There are three categories in this regard:

1. Both electrophoresis and electroosmosis are in the same direction. This normally occurs when cations are being separated. In this case, decreasing the EOF will enhance resolution at the expense of run time. Doubling the run time produces a 41% improvement in resolution.

2. Electrophoresis and electroosmosis are in opposite directions. This occurs on bare silica capillaries when anions are separated. Decreasing the EOF will enhance run time at the expense of resolution, and vice versa.

3. Electrophoresis and electroosmosis are equal but in opposite directions. Here the resolution is infinite, but so is the separation time. However, this concept was used to generate ultrahigh theoretical plate numbers (28).

It is clear that improvements in resolution are best addressed by adjustments to AjUgp, the difference in mobility between the two most closely eluting solutes in a separation. Since A/i p falls outside of the square root sign of the resolution equation, the improvement in resolution is directly proportional to the change in mobilities. This subject forms the basis for many of the chapters in this book.

2.6 JOULE HEATING

The conduction of electric current through an electrolytic solution generates heat via frictional collisions between migrating ions and buffer molecules. Since high field strengths are employed in HPCE, ohmic or Joule heating can be substantial. There are two problems that can result from Joule heating:

1. Temperature changes due to ineffective heat dissipation 2. Development of thermal gradients across the capillary

If heat is not dissipated at a rate equal to its production, the temperature inside the capillary will rise and eventually the buffer solution will outgas. Even a small bubble inside of the capillary disrupts the electrical circuit. At moderate field strengths, outgassing is not usually a problem, even for capillaries that are passively cooled.

The rate of heat production inside the capillary can be estimated by

^ = - ^ , (2.18) dT LA

where L = capillary length and A = the cross-sectional area. Rearranging this equation using J = V/R, where the resistance R = L/kA and k = the conductivity.


dT

kV' (2.19)

The amount of heat that must be removed is proportional to the conductivity of the buffer, as well as the square of the field strength.

Lacking catastrophic failure (bubble formation), the problem of thermal gradients across the capillary can result in substantial band broadening (29-31). This problem is illustrated in Figure 2.13. The second law of thermodynamics states heat flows from warmer to cooler bodies. In HPCE, the center of the capillary is hotter than the periphery. Since the viscosity of most fluids decreases with increasing temperature, Eq. (2.4) and (2.8) predict that both mobility and EOF increase as the temperature rises.

This situation becomes similar to laminar flow where the electrophoretic or electroosmotic velocity at the center of the capillary is greater than the velocity near the walls of the capillary. The temperature differential of the buffer between the middle and the wall of the capillary can be estimated from

AT = 0.24 4K

(2.20)

where W = power, r = capillary radius, and K = thermal conductivity of the buffer, capillary wall, and polyimide cladding. A 2-mm-i.d. capillary filled with 20 mM CAPS buffer draws 18 rtiA of current at 30 ky giving a AT of 75°C. A 50-|lm-i.d. capillary filled with the same buffer draws only 12 |LIA of current, yielding a AT of 50 m°C. Since the thermal gradient is proportional to the square of the capillary radius, the use of narrow capillaries facilitates high resolution. On the other hand, the use of dilute buffers or isoelectric buffers (32) permits the use of wider bore capillaries, but the loading capacity of the separation is reduced.

ep

r AT

FIGURE 2.13 Impact of the radial temperature gradient on electrophoretic and electroosmotic flow.

2.6 Joule Heating 45

The requirement for narrow-bore capillaries comes with a price due to the short optical path length. If a solution is injected equivalent to 1% of the capillary volume of a 50 cm x 50 |im i.d. capillary, the injection size is 9.8 nL. This small-volume injection coupled to a 50-|im optical path length provides for concentration limits of detection (CLOD) that are about 50 times poorer than by LC. Fortunately, through the use of stacking procedures (33) and extended path length capillaries (34), this gap has been narrowed considerably

The compromise between sensitivity and resolution is illustrated in Figures 2.14 and 2.15. Note in particular the cluster of peaks centered at a migration time of 31 min (26 min in Figure 2.15) in Figure 2.14. With the 50-|im-i.d. capillary, none of these peaks are baseline-resolved, but there is virtually no noise in the electropherogram. Separation of the same sample in a 25-|im-i.d. capillary (Figure 2.15) presents a different picture. The peaks are nearly baseline-resolved, but there is substantial noise in the output. This presents one of several compromises that must be made in HPCE. In this case, sensitivity and resolution are competing analytical goals.

0.300-!

0.262H

0.225H

0. I87H

0. 150H

0. uaH

0.075H

0.037H

0.00OH

\MJ WAAAJ L

io —J— 15 20

—~r-—1—-25 30

MINUTES

3S 40 " "1— 45

•'"••'" ! ""•"

50 '" 1 55

FIGURE 2.14 Separation of heroin impurities by MECC on a 50-|lm-i.d. capillary. Buffer: 85 mM SDS, 8.5 mM borate, 8.5 mM phosphate, 15% acetonitrile, pH 8.5; capillary: 50 cm (length to detector) X 50 (Xm i.d.; voltage: 30 kV; temperature: 50°C; detection: UV, 210 nm. Reprinted with permission from Anal. Chem., 63, 823 (1991), copyright © 1991 Am. Chem. Soc.


Even these electropherograms must be carefully interpreted. In both cases, the injection time was kept constant at 1 s. This means that the amount of material injected in the 25-|Ltm-i.d. capillary was a factor of four lower relative to the 50-|Llm-i.d. tube (Section 8.1). This contributes to the decreased signal-to-noise ratio observed when using the 25-|lm-i.d. capillary.

The problem of Joule heating depends on the capillary diameter, the field strength, and the buffer concentration. Recalling Figure 2.9 (50-|Lim-i.d. capillary), there was a slight increase in |Lieo as the field strength was increased. Figure 2.16 contains data from the same experiments, except a 100-|Lim-i.d. capillary is used. A marked departure from linearity is found at the higher buffer concentrations.

Higher concentration buffers are more conductive, draw higher currents, and produce more heat than more dilute solutions. In the 100-|im-i.d. capillary, this heat is not properly dissipated. As a result, the internal temperature rises, reducing the viscosity of the buffer. Since Eq. (2.8), the basic expression for elec-troosmotic velocity, contains a viscosity parameter in the denominator, Vgo increases with decreasing buffer viscosity. Because the buffer viscosity depends on temperature, the capillary heat removal system plays an important role in deciding the maximum field strength, buffer concentration, and capillary diam-

0.080-1

0.07CH

0.060H

0. 050H

0. 040-H

0.030H

0. 020H

0. oiCH

0. GOOH N'wWW \^\I\KJMM ]

12 I

16 1 1

20 24 MINUTES

1 28

1 32

1 36

1" 40

FIGURE 2.15 Separation of heroin impurities by MECC on a 25-fxm-i.d. Conditions as per Figure 2.14 except for capillary diameter. Reprinted with permission from Anal. Chem., 63, 823 (1991), copyright © 1991 Am. Chem. Soc.

2.7 Optimizing the Voltage and Temperature

11

5 9 u. ^ o

E7 to ' o

j>5

47

50 100 150 E (V/cm)

200 250

FIGURE 2.16 Effect of buffer concentration and field strength (E, V/cm) on the electroosmotic flow in a 100-|lm-i.d. capillary. Buffer: phosphate at a concentration of (a) 10 mM; (b) 20 mM; (c) 50 mM. Redrawn with permission from J. Chromatogr., 516, 223 (1990), copyright © 1990 Elsevier Science Publishers.

eter that can be successfully employed. Insufficient heat removal begins a vicious cycle leading to viscosity reduction, greater current draw, and higher temperature, further reducing the viscosity.

2.7 OPTIMIZING THE VOLTAGE AND TEMPERATURE

A. OHM'S LAW PLOTS

A means of optimizing the voltage and/or the temperature despite the buffer concentration and capillary cooling system is very desirable. An Ohm's law plot provides this tool with very little experimental work (35, 36). Simply fill the capillary with buffer, set the temperature, vary the voltage, record the current, and plot the results.

Some Ohm's law plots are shown in Figures 2.17 and 2.18 for an air-cooled and water-cooled temperature control system, respectively. Whenever the graph shows a positive deviation from linearity, the heat removal capacity of the system is being exceeded. Operating on the linear portion of the curve will generally yield the highest number of theoretical plates. As a rule of thumb, it is best to keep operating currents below 100 joA. Often, separations are run in the nonlinear section to optimize speed at the expense of plates, but it is not wise to push things too far.

Lowering the temperature below ambient can be used to extend the linear range of the Ohm's law plot. This is useful when high-ionic-strength buffers are

<

UJ

O

100

80

60

40

20

A ^

•B V

•

• •

" • . c •

V *>'

' XT

^ • ^ 0

V • ^ D

X T . ^ D

•^ 0 • = - • ^ ^ D

V ^- u"

¥ ^ D I ^^o°

1 ^ ^t° ^¥ ^ 0 ° ¥^^D

l±lll 1 H _ — 1-10 20

VOLTAGE (kV) 30

FIGURE 2.17 Ohm's law plots for capillary temperature control by air circulation. (A) no control; (B) 25°C; (C) 10°C; (D) 4°C. Redrawn with permission from J. High Res. Chromatogr., 14, 200 (1991), copyright © 1991 Dr. Alfred Heuthig Publishers.

2.7 Optimizing the Voltage and Temperature 49

1 LU

OL

O

100

80

60

40

20

A '

1 "V

• • B

• V •

• • V

• • ^ C

A 0 ^ • ^ 0

• ^ 0 -^ A „ «

• A °

X -6. a • -< 0

• ^ 0

• ^ 0

• ^ 0

,^^t° • ^ a * i ^ a

«ELf 1 1 I —

10 20 VOLTAGE (kV)

30

FIGURE 2.18 Ohm's law plots for capillary temperature control by water circulation. (A) no control; (B) 25°C; (C) 10°C; (D) 4°C. Redrawn with permission from J. High Res. Chromatogr., 14, 200 (1991), copyright © 1991 Dr. Alfred Heuthig Publishers.

necessary. These concentrated buffers are particularly useful in microprepara-tive CE (Section 9.10), increasing the linear dynamic range (Section 10.4) and suppressing wall effects (Section 3.3). Increasing the temperature can also be employed to speed the separation, since both v, due to the decreased viscosity of the buffer medium

For various instruments the Ohm's law plot is an effective means of evaluating the efficiency of their capillary cooling systems. Fluid-cooled systems are

and Vep increase about 2%/K


generally more effective than air-cooled systems, since the heat capacity of most fluids exceeds that of air.

B. CONSTANT VOLTAGE OR

CONSTANT CURRENT?

Power can be applied to the system in one of two ways. The voltage can be fixed, allowing the current to float based on the resistance of the buffer. Alternatively the current can be fixed. Most published work in HPCE is in the constant-voltage mode. There has been one report that found constant current more reproducible than constant voltage (37). Until this is better understood, both modes should be studied during methods development for CZE, MECC, and CGE separations. CITP is typically performed in the constant-current mode, or else separation time becomes long. CIEF may also benefit from the constant-current mode as well, although there is no evidence published to that effect.

2.8 CAPILLARY DIAMETER AND BUFFER IONIC STRENGTH

Some very subtle effects due to Joule heating can occur when comparing separations run on capillaries with different inner diameters, or even the same capillaries run on various instruments with different capillary cooling systems. Some of these issues are illustrated in Figures 2.19 and 2.20.

The ionic strength of the buffer influences not only the EOF and jil^^, but indirectly the viscosity of the medium. More concentrated buffers have greater conductivity and generate more heat when the voltage is applied. The viscosity depends on the temperature, and so there is also a dependence on the capillary diameter. This is shown for a series of runs in 50- and 75-|Llm-i.d. capillaries (38).

With the 50-|lm capillary, the migration times lengthen as the buffer concentration is increased. Ions in solution are always surrounded by a double layer of ions of the opposite charge. The migration of these counterions is in a direction opposite to that of the solute (Figure 2.21); hence, increasing the concentration of the buffer reduces the mobility of the solute, due to increased drag caused by countermigration of the more densely packed counterions.

With the 75-|Lim capillary, the solute migration times first increase as expected, but then they decrease. This decrease is a consequence of the significant effects of Joule heating at higher buffer concentrations. Note as well the impact of buffer concentration on peak width. Sharper peak widths at the higher buffer concentrations are due to stacking (Section 8.6). The

2.8 Capillary Diameter and Buffer Ionic Strength 51

.32 50nm Capillary 0200M

^'ULlJlJLJIjLii^ liujLi 0.075M

0.050M

O.O^M

8 10 12 14 16

TIME Crnin,) FIGURE 2.19 Effect of buffer ionic strength on peptide separations in a 50-|im-i.d. capillary. Buffers: 0.025-0.200 M phosphate, pH 2.44; voltage: 30 kV; capillary: 50 cm to detector x 50 |Lim i.d.; key: (1) bradykinin; (2) angiotensin 11; (3) TRH; (4) LRHR; (5) bombesin; (6) leucine enkephalin; (7) methionine enkephalin; (8) oxytocin; (9) dynorphin. Reprinted with permission from Techniques in Protein Chemistry 11, 1991, Academic Press, 3-19, copyright © 1991 Academic Press.

Stacking effect is more evident when the 75-|im-i.d. capillary is used. Injection time was held constant for this comparison; thus, a larger injection was made on the latter capillary. Stacking is not very noticeable when small injections are made.


.45 7S[im Capillary

2B 0.125M

.27

BS

I 11 I I 11^

UuulJJLIL

.18H

lOOM

JIU

I . II I 0.075U ILJJIUL

"ililiiL! 050M

1 ? 3.4 6 *

UMJC 02511

6 8 10 12 14 16

TIME (min.) FIGURE 2.20 Effect of buffer ionic strength on peptide separations in a 75-(Xm-i.d. capillary. Conditions as per Figure 2.19 except for capillary diameter. Reprinted with permission from Techniques in Protein Chemistry U, 1991, Academic Press, 3-19, copyright © 1991 Academic Press.

2.9 OPTIMIZING THE CAPILLARY LENGTH

The efficiency of the separation (theoretical plates) is directly proportional to the capillary length (38), provided the field strength is kept constant. The

2.9 Optimizing the Capillary Length 53

FIGURE 2.21 Countermigration of a solute against its ionic atmosphere.

limitation here is available voltage. Most instruments produce a maximum of 30 kV. Once the capillary length reaches a certain point, the field strength must be reduced and no further gains in efficiency are realized (6). Based on Eq. (2.16), the electrophoretic resolution depends on the square root of the number of theoretical plates and, thus, on the square root of the capillary length (38). Increasing the capillary length beyond the limits imposed by the voltage maximum lengthens the separation time without any substantial benefits. These effects are illustrated in Figure 2.22, where the number of theoretical plates is linear with the capillary length until 30 kV is reached. Note that the resolution increase is proportional to the square root of the number of theoretical plates.

Most chemists overly rely on the length of the capillary to perform their separations. This results in lengthy separations. Since diffusion is time related, the sensitivity of the method declines as well. If more time is spent optimizing the separation chemistry, shorter capillaries can be employed, with obvious benefits.


A A A UUU

800

600

400

200

Theoretical Plates (1

Jf

OOO's)

- ^

x'^

..4 ' ' '"

... i

^p^zzz

!

. x:—

1

Resolution

4-„™ i ji *

"i

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1.5

0.6

20 30 40 50 60 70 80 90 100

Length to Detector (cm)

FIGURE 2.22 Impact of capillary length on the number of theoretical plates and resolution.

2.10 BUFFERS

A. THE ROLE OF THE BUFFER

A wide variety of buffers (Table 2.3) can be employed in CZE. The buffer is frequently called the background or carrier electrolyte. These terms are used interchangeably throughout this book. Other terms frequently employed are co-ion and counterion. A co-ion is a buffer ion of like charge compared with the solute, a counterion a buffer ion of opposite charge.

The purpose of the buffer is to provide precise pH control of the carrier electrolyte. This is important, since both mobility and electroendoosmosis are sensitive to pH changes. The buffer may also provide the ionic strength necessary for electrical continuity. Usual buffer concentrations range from 10 to 100 mM, though there are many exceptions. Dilute buffers provide the fastest separations, but the sample loading capacity is reduced.

Buffer solutions should resist pH change upon dilution and addition of small amounts of acids and bases. Concentrated buffer solutions do this well, but they

2.10 Buffers 55

Table 2.3 Buffers for HPCE

BUFFER

ZWITTERIONIC BUFFERS

Aspartate

j3-Alanine

)8-Alanine

Histidine

MES

ACES

MOPSO

BES

MOPS

TES

DIPSO

HEPES

TAPSO

HEPPSO

EPPS

POPSO

DEB

Tricine

Glygly

Bicine

TAPS

CHES

CAPS

pKa

L99

3.55

10.24

9.34

6.13

6.75

6.79

7.16

7.2

7.45

7.5

7.51

7.58

7.9

7.9

7.9

7.91

8.05

8.2

8.25

8.4

9.55

10.4

CONVENTIONAL (Nonzwitterionic) BUFFERS

Citrate

Formate

Acetate

Lactate

Phosphate

Borate

Creatinine

3.12,

3.75

4.76

3.85

2.14,

9.14

4.89

4.76, 6.40

7.10, 13.3

MobiUty^

+31.6

+36.7

-30.8

+29.6

-26.8

-31.3

-23.8

-24.0

-24.4

-22.4

-21.8

-22.0

-26.2

-28.7 (low pKg value)

-56.6

-42.4

-35.8

-35.1 (low pKg value)

-40.0 (estimate)

+33.1

Data from J. Chromatogr., 1991; 545:391. ^Effective mobility for fully ionized buffers at 25°C (10~^ c m W s ) .

are often too conductive for use in HPCE. The buffering capacity of a weak acid or weak base is limited to ±1 pH unit of its pK . Operation outside of that range requires frequent buffer replacement to avoid pH changes (39). The buffer


should have a low temperature coefficient and not absorb significantly in the UV region, where detection occurs. Table 2.4 presents some of these data for a few common buffers.

B. BUFFER SELECTION

The selection of the appropriate buffer need not be difficult. For acids, start with borate buffer, pH 9.3, and for bases, phosphate buffer pH 2.5. These two buffer systems along with the appropriate additives will work well for most applications. If bases are not soluble in phosphate buffer, acetate buffer pH 4 may be more effective. Higher pH values may be required for basic proteins to avoid adherence to the capillary wall.

Phosphate buffers are often used for low-pH protein separations (39-41). McCormick (41) proved that phosphate ions bind to the capillary wall, reducing the impact of protein binding to anionic silanol groups. Prewashing the capillary with pH 2.5 phosphate buffer was reported to reduce protein binding as well (42). Capillaries aggressively pretreated with phosphate buffer have also been useful in this regard (43).

Borate buffers are useful for separating carbohydrates (44-49) and catecholamines (50, 51) because of specific complexation chemistry. Unless such specific interactions are identified, buffer selection based on the desired pH is usually satisfactory. Borate buffers are used to control pH in the range 8.3-10.3.

Phosphate and borate buffers have adequate buffer capacity over a wide pH range and are useful as general purpose buffers. They may not be useful for cer-

Table 2.4 Characteristics of Some Buffer Solutions

Buffer

50 mM KH2C6H5O7 (citrate)

25 mM KH2PO4, Na2HP04

8.7 mM KH2PO4+ 3.0 mM Na2HP04

10 mM Na2B407 (borate)

50 mM Tris-HCl; 16.7 mM Trls

pH at 25°C

3.776

6.865

7.413

9.180

7.382

Dilution Value^

+0.02

+0.080

+0.07

+0.01

na

Buffer Capacity^

0.034

0.029

0.16

0.020

na

Temperature Coefficient ^

-0.0022

-0.0028

-0.0028

-0.0082

-0.026

^pH change upon 50% dilution. ^pH change upon mixing 1 L of buffer with 1 gram equivalent of strong acid or base. ^Change in pH per °C. Data from pH Measurements, 1978, Academic Press.

2.10 Buffers 5 7

tain protein separations, particularly if biological activity must be maintained. These buffers also lack buffer capacity around pH 4, where acetate is quite good.

The selection of the buffer cation also plays a role in buffer conductivity (4). The correlation of the atomic radii and mobility was discussed in Section 2.1. In this regard, lithium and sodium salts are best used, since they contribute least to buffer conductivity. Dual buffering systems with lower mobility counterions (Tris-phosphate, Tris-borate, or Tris-Tricine) are used to reduce heating problems in the slab gel and can also be used in HPCE. Buffers used in the slab gel must be unreactive with Commassie or silver stain. This is not a guarantee that they will not absorb in the UV. The UV spectrum of a buffer should be checked prior to use to avoid detection problems. In capillary isoelectric focusing this problem is more acute, as ampholytes can absorb even at 280 nm.

Aromatic buffer constituents such as phthalates should be avoided because of their strong UV absorption characteristics, unless indirect detection is planned (Section 3.6). Strongly absorbing components such as carrier ampholytes (Chapter 5) prevent the use of the low-UV region for detection.

Zwitterionic buffers such as bicine, Tricine, CAPS, MES, and Tris are also used, particularly for protein and peptide separations. They are all amines, and some buffers such as Mes, Tricine, and glycylglycine bind calcium, manganese, copper, and magnesium ions. PIPES, HEPES, and Tris do not bind any of these metals, whereas BES binds only copper (52). Metal binding may be useful, or it may interfere with subsequent separations.

The advantage of the zwitterionic buffer is low conductivity when the buffer is adjusted to its pi. There is little buffer capacity when pK^ and pi are separated by more than 2 pH units. When the pi and pK^ are close together, the buffer is known as an isoelectric buffer (32). The advantage here is low current draw and reduced Joule heating, allowing higher buffer concentrations to be used. However, isoelectric buffers are poor stacking buffers because of their extremely low conductivity.

Table 2.3 provides data describing the mobility of the fully ionized buffer component. Mobility matching between the buffer and solute is used to improve the peak symmetry when the solute concentration is high relative to the buffer concentration (53). This problem, known as electromigration dispersion or simply electrodispersion (Section 2.14), is particularly troublesome when indirect detection is employed, because the buffer concentration must be low. A software program (Buffer Workshop, Scientific Resources Inc., Eastontown, NJ) is useful to help calculate buffer mobilities at any pH.

C. BUFFER PREPARATION

Titration of a buffer to the appropriate pH has some operational subtleties. In the trivial but ideal case, equimolar solutions of two different salts of the identical anion are blended to the appropriate pH. For example, to prepare a 50 mM,


pH 7 phosphate buffer, titrate a 50 mM disodium salt with 50 mM phosphoric acid. Under all possibilities, the final phosphate concentration must be 50 mM and the ionic strength must be consistent. For very critical separations, it is best to prepare buffers in large batches to reduce batch-to-batch variation. When this is done, pay attention to buffer stability and the potential for microbial growth.

In other cases, the buffer is often titrated with acid or base to adjust the pH. Under these conditions, both pH and ionic strength are being adjusted (54). Unusual effects, such as the reduction of EOF with increasing pH, have been observed that are attributable to this problem. Selecting a buffer that requires no titration or only a minor titration will minimize these ionic strength effects. In any event, it is important to exactly specify the buffer preparation in methodology. All buffers should be filtered before use through 0.21-|Llm filters.

Prepared buffers are available from many instrument manufacturers and suppliers. These solutions are manufactured in large batches, prefiltered, and put through quality control. Common recipes containing phosphate and borate along with specialty preparations for application-based methods are readily available. For small laboratories without water purification systems, water for HPCE can be purchased as well. Since reagent usage in capillary electrophoresis is minimal, the costs are low

2.11 TEMPERATURE EFFECTS

The impact of temperature on a series of peptides is illustrated in Figure 2.23. As the temperature is increased, the migration time always decreases because of the reduced viscosity of the BGE. The current increases as well due to this effect. The viscosity is, of course, inversely proportional to the temperature.

There are no significant changes in selectivity as the temperature is increased for these peptide separations. This will not always hold true, particularly when secondary equilibrium is employed. When proteins are being separated, temperature can have profound effects on the separation if unfolding occurs. This is illustrated in Figure 2.24 for a-lactoglobulin (55). At 20°C, a single peak is found at 4.5 min. As the temperature is increased, the band broadens until, finally, a sharp peak is found when the temperature reaches 50°C. The peak at 20°C represents the native form of the protein. At the intermediate temperature, multiple forms of the protein are found as unfolding begins. At 50°C, a peak representing the unfolded protein is found. Note that the migration times always decrease as the temperature is raised due to the aforementioned viscosity effects.

Elevated temperatures are frequently used during methods development to speed separations without having to cut the capillary. When secondary equilibrium is employed, particularly for chiral separations, subambient temperatures are often used.

59

-jL. Jl—LJuL....li I

15 25 35 45 55 Temperatiire (C)

4 6 8 10 n 14 " 15 25 35 45 55 Time ( Minutes ) Temperature (C)

FIGURE 2.23 Effect of temperature on time, current, and viscosity. Buffer: 50 mM phosphate, pH 2.5; voltage: 20 kV; capillary: 50 cm x 75 )im i.d. Reprinted with permission from Techniques in Protein Chemistry U, 1991, Academic Press, 3-19, copyright © 1991 Academic Press.

2.12 BUFFER ADDITIVES

Reagents are often added to the buffer solution for reasons other than control-hng pH. Known as buffer additives, these materials are used for several functions:

1. To modify mobility (secondary equilibrum) 2. To modify electroosmotic flow 3. To prevent solutes from adhering to the capillary wall 4. To maintain solubility

Table 2.5 lists of some of the reagents used for these purposes. These will be discussed in further detail throughout this text.


1129

mt

W

m

^ m

m

m

1 L

L-}l

' ' ^•'•••^'f"*"'"'*"""'^''"'*'"^'"lB"''l''*'T"""*'^*^"*i'"T"t''T™"f¥'''^P'^*^ i^ , 1

0 t 2 ^ 3 . * 5 « Tmetmin)

FIGURE 2.24 Influence of temperature on the electrophoretic behavior of cc-lactoglobulin. BGE: 100 mM borate, pH 8.3; voltage: 350 V/cm; capillary: 50 cm bare siUca. Reprinted with permission from Anal. Chan., 63, 1346 (1991), copyright © 1991 Am. Chem. Soc.

2.13 CAPILLARIES

A. FUSED SILICA

Fused-silica, polyimide-coated capillaries, similar to those used in capillary gas chromatography, are the material of choice for HPCE (Figure 2.25). Internal diameters ranging from 25 to 100 |lm are usually employed. Capillaries covering the range of 2 - to 700 |Lim i.d. are commercially available.^ Between 10- and 75-|im i.d., the i.d. tolerance is 1 |Lim. Fused silica is a good (though not ideal) material due to its UV transparency, durability (when polyimide coated), and zeta potential. The variation of the surface charge along with solute-capillary wall interaction are the most important limitations of the material. Functional-ized capillaries and buffer additives are used to overcome this limitation. Gel-filled capillaries (Chapter 6) for CGE are also commercially available.

Most instrument manufacturers employ a cartridge to contain the capillary This allows for more optimal integration with the capillary cooling system. The

^Polymicro Technologies, Phoenix, AZ 85017.

2.13 Capillaries 61

Table 2.5 Buffer Additives

Purpose Reagent Mechanism

To modify mobility

To modify EOF

To reduce wall effects

To maintain solubility

Transition metals

Cyclodextrins

Surfactants

Organic solvents

Sulfonic acids

Quaternary amines

Borate

Chelating agents

Crown ethers

Macrocyclic antibiotics

Calixarenes

Dendrimers

Cationic surfactant

Organic solvents

Linear polymers

Zwitterionic surfactant

Cationic surfactant

Polyamines

Linear polymers

Zwitterionic surfactant

Organic solvents

Urea

Complex formation

Inclusion comple

Micelle interaction

Solvation

Ion-pair formation

Ion-pair formation

Complex with carbohydrates, diols

Complex formation with metals

Inclusion complex

Inclusion complex

Inclusion complex

Inclusion complex

Dynamic coating, EOF reversal

Affects viscosity

Dynamic coating

Dynamic coating

Dynamic coating, EOF reversal

Covers silanols

Dynamic coating

Dynamic coating

Hydrophobicity

"Iceberg effect"

alignment between the capillary detection window and the detector optics is simple when a cartridge is used.

Capillaries can be purchased from the instrument manufacturers for $35-60 per capillary. The detection window is in place and the outlet side may be pre-measured for a particular instrument. Polyimide is removed at the inlet and outlet side to prevent shards of polyimide from clinging to the capillary opening. Alternatively, high-quality fused-silica capillaries can be purchased in bulk at a cost of $10-20 per meter depending on quantity.

A new fused-silica capillary can be prepared and conditioned in half an hour with materials costs of under $5.00. At such a low cost, these home-cut capillaries are disposable items. Rather than attempt regeneration of a suspect capillary, it is wise to simply replace it. It is best to dedicate a capillary to a particular application.


SeOjttm

Fused Silica

Polyimide Coating

12/im 25-75/im FIGURE 2.25 Cross-sectional view of a fused-silica capillary.

B. PREPARING A FUSED-SILICA CAPILLARY

The procedure for preparing a bare silica capillary is given below. Only a ruler, a cutter (silicon wafer or ceramic cutter), a butane lighter, methanol, and a tissue are required. The method for introducing the detection window should not be used with gel-filled or surface-treated capillaries.

L Nick the polyimide coating near the edge of the capillary as squarely as possible. Pull the capillary directly apart, making sure not to pull at too much of an angle. Measure from the cut end to the desired total length of the capillary (L^), and cut again. Observe the ends of the capillary under magnification to ensure that they are cut squarely.

2. Measure the separation length of the capillary (L^) and flame a length of about 2-3 mm with the butane lighter.^ Clean the burnt polyimide with a tissue moistened with methanol. The capillary can now be inserted into the instrument, taking care not to bend the now fragile

5Heat burns off the coating without making the glass brittle. Alternatively, concentrated sulfuric acid at 130°C will remove the coating in a few seconds. This method is required when gel-filled or functionalized capillaries are used.

2.13 Capillaries 6 3

detection window.^ The ends of the capillary can be flamed as well to remove about 1 mm of coating. This prevents problems from shards of polyimide obstructing the inlet and outlet. Swelling of the polyimide due to interaction with BGE components is prevented as well.

3. Wash the capillary for 15 min each with 1 N sodium hydroxide, 0.1 N sodium hydroxide, and BGE. Change the detector-side reservoir to BGE. The system is ready to run.

The base conditioning procedure is important to ensure the surface of the capillary is fully charged. For some methods, it is necessary to regenerate this surface with 0.1 N sodium hydroxide—in extreme cases, 1 N sodium hydroxide. This regeneration procedure is often necessary if migration times change on a regular basis. Regeneration may help if the zeta potential at the capillary wall is altered. Binding of solutes or sample matrix components may be the cause of this problem. When working at low pH, a wash with O.IN hydrochloric acid is useful to reduce the EOF

C. STORING A FUSED-SILICA CAPILLARY

For overnight shutdown, simply leave BGE in the capillary and be sure that the capillary ends are immersed in BGE. For prolonged shutdown or when removing the capillary from the system, all buffer materials must be removed from the capillary. Otherwise, the BGE will clog the capillary when evaporation occurs.

1. Rinse the capillary with water for several minutes. 2. Change both buffer reservoirs to distilled water. 3. Rinse for five minutes with distilled water. 4. Empty the appropriate buffer reservoir, and draw air through the capil

lary for five minutes. 5. Remove the capillary from the instrument. 6. For capillaries not part of a cartridge assembly, slide some 0.5- to 1-mm-

i.d. Teflon tubing over the optical window and gently secure with tape or septa.

Cleaning a capillary can be done offline using a simple 1-mL syringe and some polyethylene tubing. This is particularly advantageous during methods development, since instrument time is not wasted. Sleeve some tubing over the syringe needle and capillary. It is possible to flush water and then air through

^For those not wishing to prepare a detection window, a fluorocarbon-coated capillary (CElect-UVT) is available from Supelco, Bellefonte, PA. This capillary cannot be used in with liquid cooling systems containing fluorocarbons, since the coating will become brittle.

the capillary manually. A kit for conditioning CEC capillaries, available from Unimicro Technologies, can also be used on bare silica.

D. COATED CAPILLARIES

Coated capillaries are often used to prevent proteins from adhering to the capillary wall. They are also used to reduce the EOF if the application or mode of electrophoresis requires such a reduction. Hundreds of papers describing various coatings and coating procedures have appeared in the literature. As traditional silane chemistry is used to prepare the coatings, most of these papers describe coatings that are not stable at alkaline pH.

The advantage of the coated capillary for protein separations is that it is often possible to get a good separation without using additives to the background electrolyte. The disadvantages include cost and the aforementioned problems with capillary stability.

Table 2.6 lists commercially available capillaries. The uses of some of these capillaries will be described in the appropriate sections of this book. When a capillary is purchased from a vendor, the polyamide at the point of detection is already removed. Should you need to remake a window, do not flame the capillary since that will destroy the inner coating. Instead, place a drop of concentrated sulfuric acid where the window is to be made. Then place the capillary a few inches above a butane lighter flame. The window will appear when the acid temperature reaches 130°C. Carefully rinse the acid off of the capillary and examine the window under magnification to determine clarity

2.14 SOURCES OF BAND BROADENING

In Section 2.4, Jorgenson and Lukacs's model describing the efficiency of HPCE is described. This model assumes that diffusion is the only cause of band broadening. At low voltages, molecular diffusion is a leading cause of band broadening. At higher voltages, the Joule heating problem causes parabolic flow. Adsorption of solutes at the wall also leads to dispersion, as does electromigra-tion dispersion (see below). Extracolumn processes such as injection and detection can also lead to dispersion (Sections 8.1 and 9.5).

The peak variance or dispersion can be expressed by

^ ' = ^ d i f f < j + < a l l + ^det + ^heat + êd > ( 2 - 2 1 )

where cTdiff, ofnj. âp. < et> êat. ^^d (jJd are the respective variances due to diffusion, injection, capillary, detection. Joule heating, and electromigration

2.14 Sources of B

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dispersion. Because of the additivity of the square of the variances, the greatest contributor to variance becomes the hmiting factor.

The problem can best be articulated as follows. Since

N = (2.22)

a 1,000,000-plate separation on a 50-cm capillary gives G^ = 0.25 mm^. For a 100,000-plate separation, G^ = 2.5. Thus, it is reasonably simple to maintain the efficiency for a 100,000-plate separation, but quite difficult when the plate count approaches 1,000,000.

Some of the key sources of dispersion along with their importance and remedy are given in Table 2.7. On-capillary dispersion comprises contributions from injection overload, hydrostatic flow (siphoning), diffusion, adsorption (wall effects), and Joule heating. Injection overload, electrodispersion, antistacking (high-ionic-strength sample), and hydrostatic flow (57) due to fluid imbalances are easily avoided. Siphoning is unimportant for small-diameter capillaries unless the height differential becomes extreme. The contribution from diffusion, expressed by Eq. (2.13), is directly proportional to the separation time. The faster the separation, the sharper the peak.

The variance contribution from Joule heating has been extensively studied by Hjerten (58), Foret et al (59), Grushka et al (30), Knox (31), and Jones and Grushka (29). Grushka et al (30) concluded that temperature effects are negligible in narrow-bore capillaries. Use of wide-bore capillaries is possible when

Table 2.7 Sources of Band Broadening in HPCE

Process

Antistacking

Adsorption on wall

Detection window

Diffusion

Electrodispersion

Injection size

Joule heating

Poorly cut capillary

Siphoning

Importance

++++

++++

+

+++

+++

+++++

++

++

+

Solution

Reduce injection size Increase buffer concentration Reduce ionic strength of sample

Use coated capillary or buffer additive

Reduce slit width if possible

Reduce separation time

Increase buffer concentration Reduce solute concentration Mobility match buffer to solute

Reduce injection size Use stacking buffers

Reduce voltage

Ensure capillary is cut squarely

Balance fluid level of reservoirs

2.14 Sources of Band Broadening 67

low-conductivity buffers are used. The problem with low-conductivity buffers is decreased loading capacity and increased wall effects. Using Ohm's law plots (Section 2.7) helps ensure that Joule heating is not a significant cause of dispersion. Operation at the voltage prescribed by the Ohms's law plot minimizes diffusion-related dispersion since the field strength is maximized as well.

The mathematics for some of these sources of peak dispersion is covered in the individual sections along with more experimental detail. For a thorough review of this often complicated theory, the paper written by Gas et al. (60) should be consulted.

Another form of band broadening is known as electromigration dispersion or simply electrodispersion. The appearance of triangular or saw-toothed peaks when the solute concentration is high is indicative of this phenomenon. This process, which is intrinsic to the electrophoretic process, may be observed whenever two conditions simultaneously occur:

1. The solute concentration approaches the concentration of the BGE co-ion. 2. The solute's mobility differs from the mobility of the BGE co-ion.

To totally eliminate electrodispersion, the solute concentration must be at least be 50-fold lower than the BGE concentration or the mobilities of both solute and co-ion must closely match. In most cases, it is not possible to completely eliminate the effect; however, some electrodispersion is easily tolerable. Quantitative analysis is not affected as long as sufficient resolution is designed into the separation, the integration parameters are properly set, and peak areas are employed. The dispersion problem only becomes excessive at the higher (relative to BGE) solute concentrations (Section 10.4).

Electrodispersion is usually observed when employing indirect detection for small ions. The nature of indirect detection (Section 3.6) limits the concentration of the BGE. Furthermore, electrodispersion is diffusion mediated (see below), and small ions have very high diffusion coefficients. Other times, electrodispersion may appear at the higher end of a calibration curve (Section 10.4), during micropreparative separations, or on the major component of the separation when performing trace impurity determinations. If the major-component concentration is limited to 1 mg/mL and the buffer concentration is 50 mM, electrodispersion is seldom observed.

Electrodispersion was originally reported by Mikkers et al. (61). The mechanism for electrodispersion is described in Figure 2.26 for three conditions using cations as the solutes: (1) /LL^ > IX^GE^ 0-) Mc = MBGE. ^^^ O) Mc < MBGE.

where /a is the symbol for mobility. We analyze the first case in detail. If a solute has a high mobility relative to the BGE, the electrical conductivity of the solute zone is relatively high. It follows that the resistance of the solute zone is low; therefore, the field strength expressed over the zone is low relative to the field over the BGE. As diffusion occurs over the left boundary, the field strength that the solute experiences increases. The resultant acceleration in migration velocity

68

LOW FIELD

Chapter 2 Capillary Zone Electrophoresis

UNIFORM FIELD HIGH FIELD

®® ®®

UNSTABLE BOUNDARY

®® DIFF.

STABLE BOUNDARY STABLE BOUNDARY

C<MBGE UNSTABLE BOUNDARY

TIME FIGURE 2.26 Mechanism of electromigration dispersion.

causes the ions in that portion of the zone to move ahead of the other solute ions. At the right-handed boundary, the same effect occurs, except the cations recombine or restack since they are migrating toward the cathode. In case 2, no such effects can occur—the field strengths are held constant over both the solute and the BGE, since their mobilities are equal. Similar arguments can be made for case 3. If the solute concentration is sufficiently low relative to the BGE, the impact of the solute on the field strength is negligible and symmetric peaks are obtained.

Electrodispersion can be minimized by

1. Diluting the sample 2. Increasing the BGE concentration 3. Matching the mobility of the BGE co-ion to the solute

Sample dilution is often not possible because of the concomitant loss of sensitivity It is possible to use dilution to reduce electrodispersion in conjunction with an extended path length flowcell. If there is sufficient sensitivity, sample dilution is the simplest means of reducing electrodispersion.

Increasing the BGE concentration is not possible when employing indirect detection because of the reduction in sensitivity When direct detection is

References 69

employed, this technique works well up to the point at which Joule heating becomes a problem. In some cases, the capillary diameter can be reduced, thereby permitting the use of a highly concentrated BGE.

Mobility matching is very effective when indirect detection is employed; however, it is not possible to mobility match the BGE to all solutes when simultaneous separation of multiple ions is required. In this case, some of the peaks may be skewed while others are symmetric.

Other advanced techniques have been proposed to simplify mobility matching (62, 63). Since electrodispersion is not usually a problem during the course of everyday HPCE, the reader should consult the references for more information.

REFERENCES

l.Macka, M., Andersson, P., Haddad, P. R. Changes in Electrolyte pH Due to Electrolysis during Capillary Zone Electrophoresis. Anal Chem., 1998; 70:743.

2. Zhu, T., Sun, Y., Zhang, C , Ling, D., Sun, Z. Variation of the pH of the Background Electrolyte as a Result of Electrolysis in Capillary Electrophoresis. J. High Res. Chromatogr., 1994; 17:563.

3.Bello, M. S. Electrolytic Modification of a Buffer during a Capillary Electrophoresis Run. J. Chromatogr., A, 1996; 744:81.

4.Atamna, I. Z., Metral, C. J., Muschik, G. M., Issaq, H.J. Factors That Influence Mobility, Resolution and Selectivity in Capillary Zone Electrophoresis. II. The Role of the Buffer's Cation. J. Liq. Chromatogr, 1990; 13:2517.

5.Jennings, W, Analytical Gas Chromatography. 1987, Academic Press. 6.Lukacs, K. D., Jorgenson, J. W. Capillary Zone Electrophoresis: Effect of Physical Parameters on

Separation Efficiency and Quantitation. HRC & CC, 1985; 8:407. 7.Saloman, K., Burgi, D. S., Helmer, J. C. Evaluation of Fundamental Properties of a Sihca Capil

lary Used for Capillary Electrophoresis. J. Chromatogr, 1991; 559:69. 8. van de Goor, T. A. A. M., Janssen, P S. L., van Nispen, J. W, van Zeeland, M.J. M., Everaerts, E

M. Capillary Electrophoresis of Peptides. Analysis of Adrenocorticotropic Hormone-Related Fragments. J. Chromatogr, 1991; 545:379.

9.Tsuda, T., Nomura, K., Nakagawa, G. Separation of Organic and Metal Ions by High-Voltage Capillary Electrophoresis. J. Chromatogr, 1983; 264:385.

lO.Lauer, H. H., McManigill, D. Capillary Zone Electrophoresis of Proteins in Untreated Fused Silica Tubing. Anal. Chem., 1986; 58:166.

11. Walbroehl, Y., Jorgenson, J. W. Capillary Zone Electrophoresis of Neutral Organic Molecules by Solvophobic Association with Tetraalkylammonium Ion. Anal. Chem., 1986; 58:479.

12.Ermakov, S. V, Capelli, L., Righetti, P. G. Method for Measuring Very Weak, Residual Elec-troosmotic Flow in Coated Capillaries. J. Chromatogr, A, 1996; 744:55.

13.Fujiwara, S., Honda, S. Determination of Cinnamic Acid and Its Analogues by Electrophoresis in a Fused Silica Capillary Tube. Anal. Chem., 1986; 58:1811.

14. Altria, K. D., Simpson, C. F. High Voltage Capillary Zone Electrophoresis: Operating Parameter Effects on Electroendosmotic Flows and Electrophoretic Mobilities. Chromatographia, 1987; 24:527.

15. Lambert, W. J., Middleton, D. L. pH Hystersis Effect with Silica Capillaries in Capillary Zone Electrophoresis. Anal. Chem., 1990; 62:1585.

16.Tsuda, T., Nomura, K., Nakagawa, G. Open-Tubular Microcapillary Liquid Chromatography with Electro-osmotic Flow Using a UV Detector. J. Chromatogr, 1982; 248:241.

17. VanOrman, B. B., Liversidge, G. G., Mclntire, G. L., Olefirowicz, T M., Ewing, A. G. Effects of Buffer Composition on Electroosmosis Flow in Capillary Electrophoresis. J. Microcolumn Sep., 1990; 2:176.


IS.Rasmussen, H. T., McNair, H. M. Influence of Buffer Concentration, Capillary Internal Diameter and Forced Convection on Resolution in Capillary Zone Electrophoresis. J. Chromatogr., 1990; 516:223.

19.Schwer, C , Kenndler, E. Electrophoresis in Fused-Silica Capillaries: The Influence of Organic Solvents on the Electroosmotic Velocity and the f Potential. Anal. Chem., 1991; 63:1801.

20.Fujiwara, S., Honda, S. Effect of Addition of Organic Solvent on the Separation of Positional Isomers in High-Voltage Capillary Zone Electrophoresis. Anal. Chem, 1987; 59:487.

21. Corse, J., Balchunas, A. T., Swaile, D. E, Sepaniak, M.J. Effects of Organic Mobile Phase Modifiers in Micellar Electrokinetic Capillary Chromatography. J. High Resolut. Chromatogr., 1988; 11:554.

22.Hjerten, S. High-Performance Electrophoresis: Elimination of Electroendosmosis and Solute Adsorption. J. Chromatogr, 1985; 347:191.

23.Emmer, A., Jansson, M., Roeraade, J. A New Approach to Dynamic Deactivation in Capillary Zone Electrophoresis. HRC & CC, 1991; 14:738.

24.Wiktorowicz, J. E., Colburn, J. C. Separation of Cationic Proteins via Charge Reversal in Capillary Electrophoresis. Electrophoresis, 1990; 11:769.

25.Jorgenson, J. W, Lukacs, K. D. Zone Electrophoresis in Open Tubular Glass Capillaries. Anal. Chem., 1981; 53:1298.

26.Jorgenson, J. W, Lukacs, K. D. Free-Zone Electrophoresis in Glass Capillaries. Clin. Chem., 1981; 27:1551.

27.Jorgenson, J. W, Lukacs, K. D. Zone Electrophoresis in Open-Tubular Glass Capillaries: Preliminary Data on Performance. HRC & CC, 1981; 4:230.

28. Culbertson, C. T., Jorgenson, J. W. Flow Counterbalanced Capillary Electrophoresis. Anal. Chem., 1994; 66:955.

29.Jones, A. E., Grushka, E. Nature of Temperature Gradients in Capillary Zone Electrophoresis. J. Chromatogr, 1989; 466:219.

30. Grushka, E., McCormick, R. M., Kirkland, J. J. Effect of Temperature Gradients on the Efficiency of Capillary Zone Electrophoresis Separations. Anal. Chem., 1989; 61:241.

31. Knox, J. H. Thermal Effects and Band Spreading in Capillary Electro-separations. Chro-matographia, 1988; 26:329.

32. Stoyanov, A. V, Righetti, P G. Fundamental Properties of Isoelectric Buffers for Capillary Zone Electrophoresis. J. Chromatogr, A, 1997; 790:169.

33.Burgi, D., Chien, R.-L. Optimization in Sample Stacking for High-Performance Capillary Electrophoresis. Anal. Chem., 1991; 63:2042.

34.Moring, S., Reel, R. T., van Soest, R. E. J. Optical Improvements of a Z-Shaped Cell for High-Sensitivity UV Absorption Detection in Capillary Electrophoresis. Anal. Chem., 1993; 65:3454.

35. Nelson, R. J., Paulus, A., Cohen, A. S., Guttman, A., Karger, B, L. Use of Peltier Thermoelectric Devices to Control Column Temperature in High-Performance Capillary Electrophoresis. J. Chromatogr, 1989; 480:111.

36.Kurosu, Y., Hibi, K., Sasaki, T, Saito, M. Influence of Temperature Control in Capillary Electrophoresis. HRC & CC, 1991; 14:200.

37.Kurosu, Y., Satou, Y, Shisa, Y, Iwata, T. Comparison of the Reproducibihty in Migration Times between a Constant-Current and a Constant-Voltage Mode of Operation in Capillary Zone Electrophoresis. J. Chromatogr, A, 1998; 802:391.

38. McLaughlin, G., Palmieri, R., Anderson, K., Benefits ojAutomation in the Separation ofBiomole-cules by High Performance Capillary Electrophoresis, in Techniques in Protein Chemistry II, J. J. Vil-lafranca, Ed. 1991, Academic Press: 3.

39.Tran, A. D., Park, S., Lisi, P J., Huynh, O. T, Ryall, R. R., Lane, P A. Separation of Carbohydrate-Mediated Microheterogeneity of Recombinant Human Erythropoietin by Free Solution Capillary Electrophoresis. Effects of pH, Buffer Type and Organic Modifiers. J. Chromatogr, 1991; 542:459.

40. Strickland, M., Strickland, N. Free-Solution Capillary Electrophoresis Using Phosphate Buffer and Acidic pH. American Lab, 1990; November:60.

References 71

41.McCormick, R. M. Capillary Zone Electrophoretic Separation of Peptides and Proteins Using Low pH Buffers in Modified Silica Capillaries. Anal. Chem., 1988; 60:2322.

42. Zhu, M., Rodriguez, R., Hansen, D., Wehr, T. Capillary Electrophoresis of Proteins under Alkaline Conditions. J. Chromatogr., 1990; 516:123.

43.McNerney, T. M., Watson, S. K., Sim, J.-H., Bridenbaugh, R. L. Separation of Recombinant Human Growth Hormone from Escherichia coli Cell Pellet by Capillary Zone Electrophoresis. J. Chromatogr., A, 1996; 744:223.

44. Honda, S., Iwase, S., Makino, A., Fujiwara, S. Simultaneous Determination of Reducing Monosaccharides by Capillary Zone Electrophoresis as the Borate Complexes of N-2-Pyridylgly-camines. Anal. Biochem., 1989; 176:72.

45.Hoffstetter-Kuhn, S., Paulus, A., Gassmann, E., Widmer, H. M. Influence of Borate Complexa-tion on the Electrophoretic Behavior of Carbohydrates in Capillary Electrophoresis. Anal. Chem., 1991; 63:1541.

46.Honda, S., Suzuki, S., Nose, A., Yamamoto, K., Kakehi, K. Capillary Zone Electrophoresis of Reducing Mono- and Oligo-saccharides as the Borate Complexes of Their 3-Methyl-l-phenyl-2-pyrazolin-5-one Derivatives. Carbohydrate Research, 1991; 215:193.

47.Klockow, A., Amado, R., Widman, H. M., Paulus, A. The Influence of Buffer Composition on Separation Efficiency and Resolution in Capillary Electrophoresis of 8-Aminonaphthalene-1,3,6-trisulfonic Acid Labeled Monosaccharides and Complex Carbohydrates. Electrophoresis, 1996; 17:110.

48. Plocek, J., Chmelik, J. Separation of Disaccharides as Their Borate Complexes by Capillary Electrophoresis with Indirect Detection in the Visible Range. Electrophoresis, 1997; 18:1148.

49. Rydlund, A., Dahlman, O. Efficient Capillary Zone Electrophoretic Separation of Wood-Derived Neutral and Acidic Mono- and Oligosaccharides. J. Chromatogr, A, 1996; 738:129.

50.Tanaka, S., Kaneta, T., Yoshida, H. Separation of Catecholamines by Capillary Zone Electrophoresis Using Complexation with Boric Acid. Anal. Sciences, 1990; 6:467.

51. Kaneta, T., Tanaka, S., Yoshida, H. Improvement of Resolution in the Capillary Electrophoretic Separation of Catecholamines by Complex Formation with Boric Acid and Control of Elec-troosmosis with a Cationic Surfactant. J. Chromatogr, 1991; 538:385.

52.Westcott, C. C , pH Measurements. 1978, Academic Press. 53.Sustacek, V, Foret, F, Bocek, P. Selection of the Background Electrolyte Composition with

Respect to Electromigration Dispersion and Detection of Weakly Absorbing Substances in Capillary Zone Electrophoresis. J. Chromatogr, 1991; 545:239.

54.Vindevogel, J., Sandra, P Simultaneous pH and Ionic Strength Effects and Buffer Selection in Capillary Electrophoretic Techniques. J. Chromatogr, 1991; 541:483.

55. Rush, R. S., Cohen, A. S., Karger, B. L. Influence of Column Temperature on the Electrophoretic Behavior of Myoglobin and a-Lactalbumin in High-Performance Capillary Electrophoresis. Anal. Chem., 1991; 63:1346.

56. Nashabeh, W, Rassi, Z. E. Capillary Zone Electrophoresis of Proteins with Hydrophilic Fused-Silica Capillaries. J. Chromatogr, 1991; 559:367.

57. Grushka, E. Effect of Hydrostatic Flow on the Efficiency in Capillary Electrophoresis. J. Chromatogr, 1991; 559:81.

58.Hjerten, S. Free Zone Electrophoresis. Chromatogr Rev., 1967; 9:122. 59. Foret, F, Demi, M., Bocek, P Capillary Zone Electrophoresis: Quantitative Study of the Effects

of Some Dispersive Processes on the Separation Efficiency. J. Chromatogr, 1988; 452:601. 60. Gas, B., Stedry, M., Kenndler, E. Peak Broadening in Capillary Zone Electrophoresis. Elec

trophoresis, 1997; 18:2123. 61.Mikkers, F E. P., Everaerts, F M., Verheggen, T. P E. M. Concentration Distributions in Free

Zone Electrophoresis. J. Chromatogr, 1979; 169:1. 62. Williams, R. L., Vigh, G. N-(Polyethyleneglycol Monomethyl Ether)-N-Methylmorpholinium-

Based Background Electrolytes in Capillary Electrophoresis. J. Chromatogr, A, 1997; 763:253. 63. Williams, R. L., Vigh, G. Polyethylene Glycol Monomethyl Ether Sulfate-Based Background Elec

trolytes in Capillary Electrophoresis. J. Chromatogr, A, 1996; 744:75.

CHAPTER 3 Capillary Zone Electrophoresis Methods Development

3.1 Introduction 3.2 Mobility 3.3 Solute-Wall Interactions 3.4 Separation Strategies 3.5 Secondary Equilibrium 3.6 Applications and Techniques

References

3.1 INTRODUCTION

Separations by CZE are performed in a homogeneous carrier electrolyte—the electrolyte in both reservoirs and the capillary are the same. Also known as free-solution capillary electrophoresis, CZE is further distinguished from other forms of HPCE by the absence of a gel or polymer network (Chapter 6), a pH gradient (Chapter 5), or a heterogeneous electrolyte system (Section 8.6).

The separation mechanism is illustrated in Figure 3.1. Ionic components are separated into discrete bands when each solute's individual mobility is sufficiently different from all others. Separations of small ions, small molecules, peptides, proteins, viruses, bacteria, and colloidal particles have been reported. Zwitterions such as peptides are easily separated, since the ionic charge can be fine-tuned by careful adjustment of the buffer pH.

Four fundamental features are required for good separations by CZE:

1. The individual mobilities of each solute in the sample differ from one another.

2. The background electrolyte is homogeneous and the field strength distribution is uniform throughout the length of the capillary.

73


DETECTOR •^WINDOW

DIRECTION OF SEPARATION

^ : / * /*1

FIGURE 3.1 Schematic representation of a three-component separation by CZE at the moment of injection (top) and after separation (bottom).

3. Neither solutes nor sample matrix elements interact or bind to the capillary wall.

4. The conductivity of the BGE exceeds the total conductivity of the sample components, unless very small injections are made.

3.2 MOBILITY

The fundamental parameter governing CZE is electrophoretic mobility (Eq. 2.4). Since mobility depends on a solute's charge/size ratio, the buffer pH is the most important experimental variable. The impact of pH on mobility, corrected for EOF, is illustrated in Figure 3.2 (1). This illustration is known as a mobility plot. The starting point for separations is pH 2.5 phosphate buffer for bases and pH 9.3 borate buffer for acids, and the mobility plot can be used to fine-tune separations. More importantly, the mobility plot determines whether the optimal pH is selected to yield the most rugged and reproducible method. For exam-

3.2 Mobility

MOBIUTY i(r*cirfWs

6n

75

FIGURE 3.2 Effect of buffer pH on electrophoretic mobility. Reprinted with permission from J. Chromatogr., 480, 35 (1989), copyright © 1989 Elsevier Science Publishers.

pie, at pH 4.5, glutamate and acetate will separate. It is clear from the mobility plot that pH 7 will yield a more rugged separation, since small variations in buffer pH will still permit the separation to occur.

Among the solutes described in Figure 3.2 are acids, zwitterions, and an alkali metal, sodium. Within the pH range covered in the figure, the net charge on sodium is constant, as is the mobility. Acetate and glutamate are negatively charged and show electrophoretic mobility toward the positive electrode (negative mobility). Zwitterions like amino acids, proteins, and peptides show charge reversal at their pi concurrent with shifts in the direction of electrophoretic mobility.


A mobility plot is an invaluable tool for methods development. At a glance, the optimal pH for separation is clear and problem areas exhibiting poor separation are obvious. When poor separations are noted at any pH, secondary equilibrium (Section 3.5 and Chapter 4) can be employed to improve the separation. The pJ or pK^ of a solute can frequently be estimated from the mobility plot, though careful control of ionic strength is required for accurate determinations (2, 3).

The net charge on a solute at any pH can be calculated using the Henderson-Hasselbalch equation for bases,

pH = pK, +log - ^ , (3.1)

and for acids,

pK, = pH + log ( 1 ^

, a - l . (3.2)

where a = the fraction ionized. For monovalent ions, the calculation is straightforward. For zwitterions such as amino acids, the contributions of the acidic and basic portions, and side chains if any, must be combined to calculate the net charge as shown in Table 3.1. In Figure 3.3, the Henderson-Hasselbalch equation is solved for bases between pH 2.5 and 11.0. By definition, a base is 90% charged when the pH is one unit below the pK^ and 10% charged when the pH is one unit above the pK . When the pH is two or more units greater or less than the pK , assume that the basic solute is neutral or fully charged, respectively. The opposite relations hold true for acidic substances.

The net charge on small peptides or any polyvalent molecule is calculated by combining the net charge from all ionizable moieties. For peptides, the pK^ values for the C and N terminus along with values for side chains must be used to determine the contribution to the charge. For proteins and large peptides, this simple model does not work because of charge suppression; more complex calculations are required (4, 5).

Table 3.1 Calculation of net charge for Lysine at pH 7.

H2N—CH— 1

1 1 NH2

COOH

Functional Group

Carboxylate

a-Amino group

£ -Amino group

pKa

2.18

8.95

10.53

Net Charge

Charge

-1.000

+0.989

+1.000

+0.989

3.2 Mobility

Fraction Ionized

77

1

0.8

0.6

0.4

0.2

^ PXaV

^ i L - ^

4\

1 1^ .

5\

j!>"-r±r

6 \

^rr™«Llrr"t:^

. 7

••r^r«-Il!::r:a-

\ 8 ^

-rrrff^illrZa

\ 9

':s5s*«lll::zi±s»s^^ 3 4 5 6 7 8 9 10 11

pH

FIGURE 3.3 Solution for the Henderson-Hasselbalch equation for bases of specified pK from pH 2.5 to 11.

These calculations can be useful to correlate mobility with pH. Grossman et al. (6) developed an empirical relationship that linked mobility to a complex function, ln(q + l)/nO^^, where q is the charge and n is the number of amino acids. Deyl et al. (7, 8) and Rickard et al. (9) found a best fit correlation for mobility with q/M^'^, where M is the molecular weight. This is consistent with Offord's model, which describes mobility of large molecules. For small molecules, Ml/3 provides a better fit. Grossman's model (6) falls between these two values, as might be expected when separating peptides containing between 3 and 39 amino acids. The accuracy of the q/MW / versus mobility model is illustrated in Figure 3.4 (9); in the figure, data from a series of peptides from two separate digests separated by CZE at three different pH values are plotted.

If the pKa and molecular weight of a substance are known, the use of mobility calculations to select the initial experimental conditions can be a worthwhile undertaking. Although optimal separation conditions cannot be predicted using this model, the calculations are effective as a first approximation.

The profound effect of pH on mobility is illustrated in Figure 3.5 for two peptides differing by one amino acid with sequences AFKAING and AFKADNG (10). At pH 2.5, the calculated charges on these two peptides are 1.41 and 1.36, respectively. At pH 4.0, the calculated charges become 1.02 and 0.46. It is expected and observed that greater resolution is found for the higher pH buffer.


o S

o o n CL O w

O

UJ

3.000e-4'

1.000e-4'

-1.000e-4

.!)noOA^-

. ^ y^^CsL

• * • 1 '

'V' no a

» 1

^

o

[ »

a ,

» o

o > « ^

a

1 ' '»' » " 1 " »' • 1 » ' '1

-0.04 -0.02 -0.00 0.02 0.04 o.oc

FIGURE 3.4 Correlation of mobility and q/MW^^ for a human growth hormone digest (separated at pH 2.35, 8.0, and 8.15) insulin-like growth factor 11 digest (separated at pH 2.35 and 8.15). Reprinted with permission from Ana/. Biochem., 197,197 (1991), copyright © 1991, Academic Press.

since the mobilities are better distinguished. While one would expect longer migration times as the pH is increased (the charge on the peptides is less positive) , the increase in the EOF partially negates this effect.

During the course of methods development, peak broadening and/or peak tailing may be noted. This may be due the adherence of the solute to the capillary wall. If this occurs, the wall effects must be eliminated before any meaningful methods development can be completed.

3.3 SOLUTE-WALL INTERACTIONS

A. THE PROBLEM OF WALL EFFECTS

A key advantage of HPCE compared with HPLC is the absence of the chromatographic packing. The vast surface area of the packing material is responsible in part for irreversible adsorption of solutes, particularly proteins. The composition of the capillary surface, though, still provides opportunities for protein adsorption. Binding of solutes to the capillary wall leads to band broadening, tailing, and irreproducibility of separations. If the kinetics of adsorp-tion/desorption are slow, broadened tailed peaks occur. Irreversible adsorption leads to modification of the capillary, altered EOF, and loss of resolution.

3.3 Solute-Wall Interactions 79

\ ^

10 TIME (min.)

10

FIGURE 3.5 Effect of buffer pH on the selectivity of peptide separations by CZE. Capillary length: 45 cm to detector (65 cm total) x 50 [im i.d.; BGE: citric acid, 20 mM, (A) pH 2.5, (B) pH 4.0; field strength: 277 V/cm; current: in A, 24 |lA, in B, 12 |lA; temperature: 30°C; detection: UV, 200 nm; peptides: (1) AFKAING, (2) AFKADNG. Reprinted with permission from Anal Chan., 61, 1186 (1989), copyright © 1989 Am. Chem. Soc.

Figure 3.6 illustrates the electrostatic binding of a protein to the capillary wall. At most pH values, the capillary wall has a negative charge due to silanol ionization. Separation of a protein at a pH below its pJ produces a cationic solute that ion-pairs to the capillary wall. Hydrophobic binding may occur as well between the epoxide moiety of fused silica and a hydrophobic solute. Since most separations occur in aqueous media, hydrophobic solutes are not well solvated, further enhancing this potential binding mechanism.


•MUMUttUiiriiiiiUiuifaUriiyd^^

FIGURE 3.6 Illustration depicting the ion-pair formation between a positively charged protein and the negatively charged capillary wall.

The problem of wall binding is most severe for large molecules. This is easily understood from the illustration in Figure 3.7. A small molecule can have but a single point of attachment to the capillary wall. A large molecule can lie

LARGE MOLECULE SMALL MOLECULE FIGURE 3.7 Ion-pair formation between large molecules results in multiple points of attachment with the capillary wall. This is not possible for small molecules.


down on the wall and ion-pair in many places. It then becomes for difficult for the large molecule to dislodge from the wall, since all points of attachment must simultaneously be broken. For small molecules, wall effects usually result in Gaussian band broadening and the effect is slight. This has been studied for lan-thanide ions (11), where a coated capillary proved more efficient.

Adsorption or retention in HPCE is determined by a solute's adsorption/des-orption kinetics with the capillary wall. A first approximation of the impact of wall effects can be understood using a random walk model from chromatographic theory (12). Random walk theory considers a solute moving down the capillary in discrete steps. The peak variance is expressed as

^s = 2 ( - ^ ) ^ v , p t , L , (3.3) 1 + fe

where t^ = the time for adsorption. Retention occurs whenever t , the time for desorption, is greater than t , and

by definition, t^ = tjk\ The time for adsorption, t , is a function of the diffusion coefficient, the capillary diameter, and the probability of binding to the capillary wall. Table 3.2 contains data showing the molecular weight and the diffusion coefficient for various molecules. The kinetics of mass transport to the capillary wall are slower for large molecules, and this in part indicates wall effects are more severe as the molecule weight increases.

When solutes adhere to the wall, peak tailing may be observed since a des-orbed solute does not return at once to the buffer solution. Retained solutes have a migration velocity of zero. Solutes in the buffer move at a rate determined by their migration velocity and, thus, move ahead of retained material. If we

Table 3.2 Diffusion Coefficients of Large and Small Molecules

Compound

j3-alanine

Phenol

Citric acid

Cytochrome c

^-Lactoglobulin

Catalase

Myosin

Tobacco mosaic virus

Molecular Weight

89

94

192

13,370

37,100

247,500

524,800

590,000

D(cmV X 105)

0.933^

0.84^

0.66P

0.114^

0.075'

0.04F

O.OIF

0.0046^

^From Handbook of Physics and Chemistry, 46th ed., 1965, CRC, p. F46. ^From, B. L. Karger, L. R. Snyder and C. Horvath, An Introduction to Separation Science, 1973, John Wiley & Sons, p. 79. ''From A. L. Lehninger, Biochemistry, 1970, Worth Publishers, pp. 136-137.

solve Eq. (3.3) for the variance and plot the decrease in the number of theoretical plates versus k' (Figure 3.8), the dramatic impact of wall effects on efficiency is apparent. To achieve the theoretical efficiency of CZE, k' must approach zero. As the figure illustrates, even modest retention will lead to severe band broadening. In the worst case scenario, no elution occurs—the solute is completely bound to the capillary wall. This simple random walk model only estimates the impact of wall effects on efficiency. More sophisticated calculations have appeared in the hterature in 1995 (13, 14). In any event, the appropriate buffer additives or capillary coatings are required to minimize this form of band broadening.

Using a clever experimental apparatus with multiple detectors, Towns and Regnier (15) were able to measure the binding of proteins to the capillary wall. Some of their data are reproduced in Table 3.3. Under their experimental conditions, all proteins showed some binding. As expected, the high-pl proteins bound most strongly, owing to their positive charge at pH 7.

Wall effects on bare sihca have proved to be a difficult problem since the early days of HPCE (16). Since then, several solutions have been proposed including the use of

1. Extreme pH buffers 2. High-concentration buffers 3. Amine modifiers

-J r\ a (fl -I O < -o — h- — UJ 5

FIGURE 3.8 Impact of wall retention on efficiency where Vep = 1 mm/s, L = 500 mm, and ta = 100 ms (0), 50 ms (+), 10 ms (D).

Lysozyme

Cytochrome c

Ribonuclease A

Chymotrypsinogen

Myoglobin

Conalbumin

Carbonic anhydrase

/3-Lactoglobulin B

j8-Lactoglobulin A

Ovalbumin

Pepsin

11.1

10.2

9.3

9.2

7.3

6.3

6.2

5.2

5.1

4.7

3.2


Table 3.3 Percent Recovery for Proteins of Varying pi on an Uncoated Capillary

Protein pi Percent Recovery

0

0

0

0

63

66

72

74

76

81

90

Determined using two detectors 60 cm apart on a 75-|Xm-i.d. capillary. Conditions: 10 mM phosphoric acid, pH 7; detection at 214 nm; 300 V/cm, 30 lA.

Data from reference (15).

4. Dynamically coated capillaries 5. Treated or functionalized capillaries

B. EXTREME P H BUFFERS

In 1986, Lauer and McManigill (17) reasoned that anionic proteins should be repelled from the anionic capillary wall. Selecting a pH 1-2 units above the pJ of a protein produces separations free of wall effects. There are several limitations to this approach.

1. Hydrophobic binding is not eliminated. 2. Proteins with high pi vlaues, such as histones, do not form anions until

pH 13 is reached. To reach that pH, 100 mM sodium hydroxide is required, a solution that is too conductive to be a useful electrolyte.

3. Alkaline pH values may not be optimal for the separation.

Despite these limitations, this useful approach was the first to employ charge manipulation to reduce wall interactions.

Two years later, McCormick (18) chose a different approach. At pH 1.5, the silanol groups at the capillary wall are not ionized. While the proteins are cationic at that pH, electrostatic interactions should be eliminated. McCormick


proposed that phosphate groups bind to the capillary wall, further decreasing the activity of the silanol groups. As with working at a high pH, low-pH separations may not be ideal. This is particularly important when separating similar proteins where the charge to mass ratios are all similar at pH extremes.

C. HIGH-CONCENTRATION BUFFERS

Green and Jorgenson (19) found that buffers with high concentrations of salts decreased adsorption of proteins on the capillary wall. The added salts probably occupy potential adsorption sites. Potassium sulfate was the additive of choice based on performance and UV absorption background. A buffer comprising 100 mM CHES, pH 9, with 250 mM potassium sulfate and 1 mM EDTA produced 140,000 theoretical plates for trypsinogen. The approach was limited, because low field strengths and narrow-i.d. capillaries were necessary due to Joule heating problems. A combination of low field strength and low EOF yielded prolonged run times. Sensitivity was also problematic, because of background UV absorption by the electrolyte.

To address the limitations of high-ionic-strength buffers, Bushey and Jorgenson (20) substituted a zwitterionic species such as glycine, betaine, or sar-cosine for the ionic salt. When the buffer pH is adjusted to the pi of the zwitterion, its net charge, mobility, and conductivity approach zero. The best results were found using 40 mM phosphate/250 mM potassium sulfate, pH 7; 100 mM CHES/250 mM potassium sulfate, pH 9; and 40 mM phosphate/2 M betaine/100 mM potassium sulfate, pH 7.6. In principle, any zwitterionic buffer could be used at the appropriate pH provided the UV absorption is sufficiently low.

Even modest increases in the ionic strength can have profound impact on the separation. Increasing the buffer concentration from 10 mM Tricine, pH 8.1, to 100 mM Tricine at the same pH gave dramatic improvements for the separation of a human growth hormone tryptic digest (21).

D. AMINE MODIFIERS

Amine derivatives such as triethylamine have been used for years to suppress silanol effects in HPLC. Alkyldimethylamines have been shown to be most effective (22). In 1986, Lauer and McManigill (17) added 5 mM putrescine (1,4-diaminobutane) to improve the resolution and peak shape of myoglobin. Since that time, many papers have appeared reporting the employment of polyamines as electrolyte additives to reduce both EOF and protein binding to the capillary wall (23-30). Used at concentrations as low as 1 mM or as high as 30-60 mM, 1,3-diaminopropane improves resolution of basic proteins such as lysozyme.


cytochrome c, and ribonuclease. Monovalent amines such as triethylamine or n-propylamine were not as effective. The use of 1,3-diaminopropane allows separations of basic proteins at pH values below their isoelectric points. Since the pH can be independently controlled over a wide range, a buffer can be optimized without considering wall effects. Separations are shown in Figure 3.9. The problem with 1,3-diaminopropane is volatihty and toxicity As a result, 1,4-diaminobutane (27-29) or 1,5-diaminopentane (26) have been used in its place.

A) pH 9 . 0 H 6

iLXl

C) pH 7 .0

H

M l M V . •^w-^

20 Time (min)

40

b) pH 6 . 2

j^jpKiijf

0* 20' 40 Time (min)

1 3

2

E) pH 4:7

IJUL 20

Time (min) 40

f ) pH 3 .5 2 4 '^ 3

20' Time {min)

40

FIGURE 3.9 Separation of basic proteins on an untreated fused-silica capillary with diamino-propane as a buffer additive. Capillary: 75 cm (55 cm to detector) x 50 |Lim i.d.; BGE: pH values as noted in the figure with 30-60 mM diaminopropane as an additive; field strength: 200-240 V/cm; solutes: (1) lysozyme; (2) cytochrome c; (3) ribonuclease; (4) a-chymotrypsin; (5) trypsino-gen; (6) rhulL-4. Reprinted with permission from J. Microcolumn. Sep., 3, 241 (1991), copyright © 1991 Microseparations, Inc.


Another polyamine, N,N,N,N-tetramethyl-l,3-butanediamine, used between pH 4.0 and 6.5 has been effective for the separation of basic proteins (31).

E. DYNAMICALLY COATED CAPILLARIES

In 1990, Wiktorowicz and Colburn (32) reported on an approach that employs a cationic surfactant to reverse the charge on the capillary wall. Since then, numerous papers have appeared reporting on the use of that approach (33-40).

The mechanism of charge reversal is illustrated in Figure 3.10. Ion-pair formation between the cationic head group of the surfactant and the anionic silanol group occurs. The hydrophobic surfactant tail extending into the bulk solution cannot be solvated by water. Its solvation need is satisfied by binding to the tail of another surfactant molecule. As a result, the cationic head group of the second surfactant molecule is in contact with the bulk solution. The capillary wall behaves with cationic character because of this treatment, and the EOF is directed toward the positive electrode. For most separations, it is necessary to operate the HPCE instrument with reverse (sample side negative) polarity. Using this approach, a buffer pH is selected that is below the pi of the target protein. The cationic protein is now repelled from the cationic wall.

In the Wiktorowicz approach, the cationic surfactant is not present in the run buffer. The capillary is coated before the separation step. Excess surfactant is flushed from the capillary and replaced with run buffer. This coating is stable for many runs, after which the capillary is simply recoated. The coating is sufficiently stable to permit CE/MS of proteins without detection of the surfactant (39).

CAPILLARY WALL

\ / ( r\ \'V( (I

w ^s

)i k ® « ® ^

EOF

FIGURE 3.10 Pictorial representation of cationic-surfactant-mediated charge reversal at the capillary wall with concomitant reversal of the electroosmotic flow.


In Emmer's method, a fluorinated surfactant^ is added to the run buffer at a concentration of 100 jig/mL. A value of more than 300,000 theoretical plates was reported for lysozyme, a protein with a pi of 11. This approach, along with Wiktorowicz's, does not require a covalently treated capillary, extreme pH, amine additives, nor high-ionic-strength buffers.

Also of note is the use of 1 mM hexamethonium bromide or 300 |im decamethonium bromide as a dynamic coating for the resolution of protein gly-coforms (41). At such low concentrations, the charge on the capillary wall is not reversed, and so normal polarity voltage is used.

A new series of reagents^ have been shown to dramatically stabilize the EOF, resulting in highly reproducible migration times. Figure 3.11 illustrates the mechanism for this dynamic coating. First, the capillary is treated as usual with 0.1 N sodium hydroxide followed by a rinse with a polycation solution. Then, a second layer consisting of a polyanion in a buffer of the desired pH is flushed through the capillary. Figure 3.12 shows four replicate runs for a chiral separation of epinephrine. The runs are virtually superimposible, showing reproducibility seldom found in HPCE. Since the polyanion coating provides a high EOF at pH 2.5, the capillary length must be longer that for bare silica when separating bases. The reagents have been shown to work for bases at a pH below the pKg. Other applications have not been verified as of January 1999.

F. FUNCTIONALIZED CAPILLARIES

There is no shortage of publications covering the use of coated capillaries, as this sampling of references indicates (33, 34, 42-81). Unfortunately, the quest for the ideal surface for HPCE remains elusive. The preparation of capillaries with traditional silane chemistry renders the coating labile to alkaline hydrolysis. The ideal surface would be hydrophilic, stable, and exhibit no EOF or at

ipluorad FC 134 from 3M Company, St. Paul, MN. ^CElixir, Scientific Resources, Inc., Eatontown, NJ.

CAPILLARY WALL

POLYCATION LAYER

POLYANION LAYER

FIGURE 3.11 Illustration of the coating process using a polycation and polyanion to stabilize the charge on the capillary wall.


3 5"

FIGURE 3.12 Four overlaid runs of the chiral separation of epinephrine using CElixir. Capillary: 72 cm X 50 |lm i.d. BF3; voltage: 30 kV; BGE: CElixir, pH 2.5, with 20 mM dimethyl-j3-cyclodex-trin; injection: 100 mbs; temperature: 20°C; detection, UV, 200 nm; epinephrine: 1 mg/mL in water.

least have controllable EOF. The goal is to prepare a stable and inert surface that provides no retention. Even a minor amount of retention will dramatically reduce efficiency due to mass transfer effects.

An ideal surface is very hydrophilic, or a buffer additive can used to maintain hydrophilic character. Since HPCE is performed in aqueous media, any hydrophobic character in the surface treatment will result in undesirable retention. Jorgenson and Lukacs (82) prepared a silylated capillary to reduce the EOF and improve the resolution of dansyl amino acids. In 1985, Hjerten produced treated capillaries using polyacrylamide or methylcellulose as the coating (42). Solute adsorption was reduced, but the polyacrylamide coating was not very stable, particularly at high buffer pH.

Very little data have been published on most of these coatings. Polyacry-lamide-treated capillaries have received the most attention because of their history of use in slab-gel electrophoresis. These surface-treated capillaries have been commercially available for some time.^ The coating has a limited lifetime, particularly in alkaline buffers, although a stabilized version has been available for some time.' Other commercially available capillaries are noted in Table 2.6.

^Bio-Rad, Richmond, CA. 4jUSIL-FC,J&W Scientific, Folsom, CA.


A fluorocarbon-polymer-coated capillary is used in conjunction with small amounts of either cationic, anionic, or neutral fluorinated surfactants. The charge on the capillary wall and thus the EOF can be controlled by the selection of surfactant and surfactant concentration. The capillary is more tolerant of alkaline pH, presumably because the added surfactant can "repair" the capillary surface.

A capillary with a permanent positive charge^ is useful for separating positively charged proteins, since they are repelled from the capillary surface. Since the EOF is now directed toward the anode, this capillary is used in the reversed-polarity mode.

A polyvinyl-alcohol-coated capillary^ for use between pH 2.5 and 9.5 has been shown useful for many protein separations. Phosphate, Tris, and 2-amino-2-methyl-l,3-propandiol are the recommended buffers. Borate should be avoided with this capillary because of surface swelling.

Lowering of the EOF occurs in most coated capillaries. Under these circumstances, it may be necessary to reverse the voltage polarity when anions are separated. Towns and Regnier found a surface coating of 30 A is required to suppress the underlying silanol effects. The dependence of EOF on buffer pH can also be reduced (83).

The use of hydrophobic coatings with surfactant additives can produce a well-coated hydrophilic surface (83). Nonionic surfactants such as Brij-35 or Tween-20 at levels of 0.01% in the buffer yield good separations of acidic (Figure 3.13a) or basic (Figure 3.13b) proteins. Approximately 240,000 theoretical plates have been obtained for myoglobin.

G. GC CAPILLARIES

Capillaries manufactured for gas chromatography (GC) can probably be substituted for coated capillaries for a very low cost per capillary. DB-1 (dimethylpolysiloxane), DB-17 (50% methyl/50% phenylpolysiloxane), DB-Wax, and others are available precut with detection windows included.-^ These capillaries can also be purchased in bulk as GC capillaries, but detection windows must be cut using either hot sulfuric or nitric acid. The use of GC capillaries purchased in bulk has not been extensively studied with HPCE, although a reference appeared in the literature in 1996 (84).

H. COATED CAPILLARIES VERSUS BUFFER ADDITIVES

Good separations of proteins are possible using either buffer additives or coated capillaries. It is impossible to predict if one method will predominate over the

5eCAP amine, Beckman Instruments, Fullerton, CA. 6PVA capillary, Hewlett-Packard, Little Falls, DE. TJ&W Scientific, Folsom, CA.


T .002 AU

i

nv^-'^ I JU IK-W**

8 12 Time (min)

16 20

FIGURE 3.13A Electropherogram of basic proteins in an alkylsilane-treated capillary with a non-ionic surfactant buffer additive. Capillary: 50 cm x 75 \im i.d. alkylsilane-treated capillary; BGE: 10 mM phosphate, pH 7 with 0.01% Brij-35; field strength: 300 V/cm; detection: 200 nm; solutes: (1) lysozyme; (2) cytochrome c; (3) ribonuclease A; (4) a-chymotrypsinogen; (5) myoglobin.

Other. From the standpoint of cost, buffer additives are clearly preferable since bare fused-silica capillaries are disposable items. On the other hand, buffer designs are more complex using additives, and the additives may interfere with mass spectrometry. For micropreparative work, removal of the additive may also be necessary. When separating small molecules, coated capillaries are seldom necessary.

3.4 SEPARATION STRATEGIES

Much of the work reported for CZE deals with proteins, peptides, and small molecules. For many of these applications, methods development is required, since each solute is unique. The approaches toward methods development for these classes of compounds are given in this section. Methodology for applications areas such as carbohydrates, small ions, peptide mapping, and others will be covered in the applications sections.

3.4 Separation Strategies 91

Time (mln)

FIGURE 3.13B Electropherogram of acidic proteins in an alkylsilane-treated capillary with a non-ionic surfactant buffer additive. Capillary: 30 cm X 75 |Lim i.d. alkylsilane-treated capillary; conditions as per part a; solutes: (1) myoglobin; (2) conalbumin; (3) transferrin; (4) p-lactoglobulin B; (5) P-lactoglobulin A; (6) ovalbumin. Reprinted with permission from Anal Chem., 63, 1126 (1991), copyright © 1991 Am. Chem. Soc.

A. SOLUBILITY

Ensure the solute is soluble in the BGE prior to the run. Even a hint of cloudiness may mean hydrophobic material is not in solution. Spikes and/or broad peaks that may not have reproducible migration times may appear in the electropherogram. It is possible that no peaks will appear at all. Though organic solvents can be used to provide solubility, do not neglect 7 M urea as an additive. Its ability to get material in solution should not be underestimated. Adding 7 M urea to 200 mM borate buffer, pH 9.2 can be used to solubilize hydrophobic membrane proteins (85). If basic compounds are insoluble in low-pH phosphate buffer, acetate buffer may be a better choice.

B. PROTEINS AND LARGE PEPTIDES

There is no single solution to developing conditions useful for separating all proteins. Given in the following is a series of practical experiments for quickly screening a variety of conditions. Detection is best at 200 nm, though 220 or 280 nm are used for some applications if the BGE absorbs in the low UV.


1. Start with 20-50 mM borate buffer, pH 9.3, or select a pH at least 2 pH units above the protein's pJ. The borate concentration can be increased to 150 mM, though a 25-|lm-i.d. capillary may be required. See Section 3.6E for a discussion of serum protein separations.

2. For basic proteins, use a coated capillary with 20-50 mM phosphate buffer, pH 2.5 or 7.

3. Prepare a BGE identical to the matrix in which the protein is dissolved, provided the matrix does not absorb in the low UV A 25-|Lim-i.d. capillary can be used for conductive electrolytes.

4. Add a cationic surfactant to the BGE. 5. Add a poly amine to the BGE. 6. Use surfactants such as SDS above the critical micelle concentration. 7. Work with high-ionic-strength buffers such as 150 mM borate buffer,

using 25-|im-i.d. capillaries to minimize heating. 8. For insoluble solutes such as membrane proteins, add 4-7 M urea to the

BGE. 9. Try ethylene glycol as an additive (86).

Table 3.4 contains capillary and electrolyte data for a variety of real applications including separating posttranslational modifications of recombinant proteins, separation of glycoforms, clinical analysis, and quality control.

C. SMALL MOLECULES AND SMALL PEPTIDES

1. Determine the wavelength for detection. If a charged solute does not absorb in the UV region, then indirect detection must be employed. If the solute is neutral and does not absorb, it is not a candidate for HPCE unless derviatization is employed.

2. Determine if the solute is soluble in water or one of the common buffers usually employed. If insoluble, organic solvents may be used—but caution should be exercised if micelles or cyclodextrins are in the BGE, since peak splitting may occur. Addition of 7 M urea to the sample and/or BGE will get most materials in solution, but this precludes the use of wavelengths below 250 nm for detection. If the solute is highly water-insoluble, then consider nonaqueous capillary electrophoresis (NACE).

3. For acids, start with 20-50 mM, pH 2.5 phosphate buffer. The more dilute buffer will give faster separations, though at the expense of loading capacity and, occasionally, resolution. For bases, start with 20-50 mM, pH 9.3 tetraborate buffer. Bare silica capillaries are usually fine until the molecule becomes sufficiently large that there can be multiple points of attachment to the capillary wall.

4. If resolution is inadequate, a mobility plot should be performed across the average of the pK^ values of the solutes.

3.4 Separation Strategies 93

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3.5 Secondary Equilibrium 95

5. If the separation is still inadequate, then secondary equilibria should be utilized. This topic will be described in detail in Section 3.5 and Chapter 4. MECC is the most common form of secondary equilibrium.

Table 3.5 contains a listing of applications and buffer recipes for a variety of solutes.

3.5 SECONDARY EQUILIBRIUM

The use of secondary equilibrium in HPCE entails the transient interaction of solutes with an added reagent. Let us assume that two cationic solutes, A and B, are inseparable at any pH but that a reagent has been identified that may interact with the solutes. Then the following equilibrium expressions can be written:

A- + R ^ A^R, (3.4)

B+ + R ^ = ^ B^R. (3.5)

If the equilibrium is pushed too far to the left, no separation can occur, since A+ and B+ are inseparable. When the reagent interacts with the solute, the mobility decreases since the neutral reagent contributes mass without charge. However, if the equilibrium is pushed too far to the right, no separation occurs, since A+R and B+R are usually inseparable. Separation only occurs when two conditions are met:

1. Kg does not equal Kb. 2. The equilibrium is not pushed to either extreme.

Wren and Rowe developed a model that they applied to chiral recognition (137-139); their model, though, is applicable to virtually all forms of secondary equilibrium:

A^ = [C](M.-A^.XK,-K,) ^3 ^

l + [C](Ki + K^) + K.K.IC?

Here [C] is the concentration of the complexation reagent, K and Kj are the respective equilibrium constants, ^^ is the mobility of the free species, and ^2 is the mobility of the complexed species.

The equation is solved and the results are plotted in Figure 3.14. Three types of behaviors are illustrated:

1. Weak binding 2. Moderate binding 3. Strong binding

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S e

0 m

fM OX)

B

0 (N

-d

a

'0

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0 c«

'0

0 § C3

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v£)

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pH

0 4

2

s s 0 in

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0

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'^ s 0 0 "S

M

PH I/) 0 p

.

0 0 P^

0

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1^

X p

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03 ^

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s 1


Mobility Difference

0.01 0.02 0.03 0.04 0.06 0.06 0.07 0.08 0.09 Concentration (M)

0.1

FIGURE 3.14 Effect of reagent concentration on the mobility differences between two solutes when secondary equilibrium is employed. The equation is solved for strong binding ( • ) , moderate bonding (+), and weak binding (*). A AK of 10% is used in all cases.

Note that the mobihty difference between two solutes is a function of the reagent concentration and the equihbrium constant. When the equihbrium is pushed to the right, the reagent concentration must be kept low; when the equilibrium is pushed to the left, the reagent concentration must be kept high. Since the equilibrium constants are not known in advance, the reagent concentration must always be carefully studied in order to optimize the method.

The next feature to consider is the charge of the reagent and solute. To separate charged solutes, the reagent can be charged or neutral. When the solute is neutral, the reagent must be charged. This holds for all reagents, whether micelles, cyclodextrins, or anything else.

Micelles and cyclodextrins are the most common reagents employed for secondary equilibrium. More than 1000 papers have appeared in the literature reporting on the use of these reagents, and Chapter 4 will be devoted to their usage, along with related reagents. The theory developed around micelles is based on chromatographic theory as opposed to the equilibrium model described above. The chromatographic theory will be presented in the next chapter.

Chelating agents, ion-pair reagents, and transition metals are frequently used to enhance separations. Metal ions such as Cu(II), Zn(II), Ca(II), and Fe(II)

3.6 Applications and Techniques 99

often coordinate to nitrogen, oxygen and sulfur. Mosher (140) separated histi-dine-containing dipeptides with Zn(ll) and Cu(ll) additives in 100 mM phosphate buffer, pH 2.5. Addition of 20-30 mM ZnS04 gave basehne resolution of DL-His-DL-His diastereomers. Other groups that can interact with metal ions include thiol, indoyl, N-terminal amino, imidazole, and j8-carboxyglutamate. Phosphorylated proteins interact with Mg(ll) and Mn(ll). Separations of calcium-binding proteins such as calmodulin, parvalbumin, and thermolysin along with zinc-binding proteins like carbonic anhydrase and thermolysin have been reported (141,142) using the respective metal ion. A 2 mM addition of calcium yielded a 15-min shift in the migration time of carbonic anhydrase.

Silver ion is useful in separating alkenes (143). Figure 3.15 shows a separation of cis and trans isomers of a proprietary compound. A small amount of SDS was added to the BGE to keep the silver from binding to the capillary wall. Without the added SDS, peak tailing was observed due to a wall effect.

Another example using secondary equilibrium coupled with adjustment of the EOF in a short capillary to give a subminute separation of nucleosides is shown in Figure 3.16(131). The EOF of the bare silica capillary is reversed with 0.2% hexadimethrin bromide in order that it is directed, like that of the anionic solutes, toward the positive electrode. To adjust selectivity, a 20 mM solution of Mg2+ is added to the BGE. Using the 7-cm short end of the capillary for the separation produces a run time of 50 s.

Chelating reagents such as hydroxyisobutyric acid can be used to modify the mobility of lanthanides and transition metals. Borate buffer is used to form complexes with carbohydrates (144-147).

Hydrophobic interaction between peptides or proteins and alkyl sulfonic acids increases the net negative charge of the solute (148). Solutes with differing affinity for the sulfonic acid are differentiated. The alkylsulfonic acid's tail binds to a hydrophobic site on the analyte, with the anionic head group extending out into the bulk solution. Increasing the negative charge is helpful in reducing wall effects. Selectivity can be altered by selecting the length of the alkyl chain; pentanesulfonic through decanesulfonic acids are good choices. Increasing the sulfonic acid concentration generally improves selectivity at the expense of Joule heating; 50 mM is sufficient for many separations.

3.6 APPLICATIONS AND TECHNIQUES

A. CAPILLARY ION ELECTROPHORESIS

Separation of small ions is traditionally performed by ion-exchange chromatography using suppressed conductivity detection. This technique, known as ion chromatography (IC), has been successfully employed for more than 20 years. The technology, though, is not without limitations. Expensive and sometimes


13.03

1.39 acslonv

'^'^^'Mmitm.

3.35

L • ^H • •am "wi" *"

FIGURE 3.15 Separation of cis/trans isomers of an alkene using silver ion. Capillary: 26 cm (20 cm to detector) x 50 jLim i.d.; BGE: 32.5 mM borate, pH 9, 2.5 mM silver nitrate, 5.0 mM SDS; voltage: 30 kV; injection: 5 s (Beckman); temperature: 40°C, detection: UV, 213 nm.

short-lived columns are required, and the mobile phase is usually strong acid or base. Pumping systems may require frequent maintenance for peak performance. Matrix effects are commonplace, and extensive sample preparation may be required. The cost per analysis may be high.

Separation and detection of small ions such as Na" , K" , Cl~, Br", N02~, transition metal ions, and lanthanide ions by CZE present unique problems. Many of these species are completely ionized between pH 2 and 12. Other ions such as borate and carbonate have pK^ values of 9.24 and 10.25, respectively; ammonia has a pKb of 4.75. Their mobilities are substantially affected by pH. Metal ions such as Li+, Na^, and K+ are fully ionized and easily separated, but many transition metal and lanthanide ions prove challenging. Since many of these solutes do not absorb light, indirect or conductivity detection must be employed.

The subject was reviewed in 1991 by Jandik et al. (149), tracing the history of the technique back to 1967, when Hjerten separated bismuth and copper in a 3-mm-i.d. rotating capillary


0.025 T

0.015 t

0.005 t

-0.005 30 40 50

migration time (s)

FIGURE 3.16 High-speed separation of nucleoside 5'-triphosphates with magnesium ion. Capillary: 47 cm (7 cm to detector) x 50 iLim i.d.; BGE: 16 mM ammonium citrate, 10 mM citric acid, pH 5, 20 mM Mg +; voltage: +25 kV (actually reversed polarity, but voltage is positive since the short end of the capillary is utilized); temperature: 25°C; injection: electrokinetic, 5 kV for 1 s; detection: UV, 254 nm; nucleotide concentration: 5 mg/L. Capillary conditioned with 0.2% hexadimethrin bromide, 0.5 min prior to each run. Reprinted with permission from J. Liq. Chromatogr. Related Tech-nol, 22, 2389 (1999), copyright © 1999 Marcel Dekker.

A comparison between IC and CZE for a series of anions is illustrated in Figure 3.17. The peak shape and time of analysis is superior by CZE. However, the sensitivity of suppressed conductivity detection is much greater than indirect photometric detection, the technique usually employed for measuring small non-UV-absorbing ions. A series of recipes for separating anions or cations is given in Table 3.6. From this table, it appears that there many types of electrolyte systems usable for a given application.


2 4

\L2/^

a.i ' ' Vi 6.0 8.6 10.6 , 12.6 14.6 Retention Time Minutes)

FIGURE 3.17 Separation of an anion standard by (A) ion chromatography and (B) capillary zone electrophoresis. In A, column: Vydac 302IC4.6; eluent: isophthahc acid, 250 jXg/mL, pH 4.6; flow rate: 2.5 mlVmin; injection size: 25 fiL; detection: 280 nm. In B, capillary: 65 cm x 75 |Xm i.d.; BGE: 2 mM borate, 40 mM boric acid, 1.8 mM chromate titrated to pH 7.8 with diethylenetetramine; voltage: 20 ky reverse polarity; detection: indirect UV, 280 nm. Key: (1) chloride; (2) nitrate; (3) chlorate; (4) nitrate; (5) sulfate; (6) thiocyanate; (7) perchlorate; (8) bromide. Reprinted with permission from J. Chromatogr, 602, 241 (1992), copyright © 1992 Elsevier Science Publishers.

1. Control of the Electroosmotic Flow

The direct of migration of cations and anions is illustrated in Figure 3.18. For cations, the electrophoretic mobility and the EOF are both directed toward the cathode. Regardless of the mobility of the small cation, the direction of migration does not change (Figure 3.18, top). The case is different for anions. Highly mobile anions such as chloride or sulfate will migrate toward the anode despite the strong EOF (Figure 3.18, middle). Less mobile anions such as hexane sulfonate are swept toward the cathode by the EOF To determine all anions in a single run, the EOF must be reversed (Figure 3.18, bottom) with a cationic surfactant such as cetyltrimethylammonium hydroxide or a protonated polyamine such as diethylenetriamine (DETA). Once the EOF is directed toward the anode, all anions migrate in that direction. The type of cationic surfactant and even mixtures of surfactants can be used to modify the selectivity of anion separations (150).


Table 3.6 Buffer recipes for indirect detection

Analyte Electrolyte Reference

Anions

Amino acids

Carbohydrates

Creatinine

Lanthanides

Metal cations

Organic acids

Pentosane Phosphates,

polyphosphates

Phosphonic acids,

alkyl

Phospholipids

Phytate

Potassium as drug counterion

Short chain fatty acids

Sodium dodecyl sulfate

Sulfates, alkyl

5 mM chromate, 0.01 mM TTAB 5 mM chromate, 0.2 mM TTAB, pH 8.2 20 mM p-aminobenzoate, 0.07 mM TTAH, pH 9.6 2.25 mM pyromellitic acid, 6.5 mM NaOH, 0.75 mM hexamethonium hydroxide, 1.6 mM triethanolamine, pH 7.7

10 mM p-amino salicyclic acid, 0.05 mM CTAB, pH 11 10 mM p-amino salicyclic acid, 20 mM a-CD, pH 11

6 mM sorbate, pH 12.2

63 mM sodium hydroxide, 12 mM riboflavin

5 mM pyridine, 3.6 mM tartaric acid, 2 mM 18C6, pH 4

10 mM creatinine-acetate, pH 4, 2 mM HIBA

8 mM nicotinamide, pH 3.2, 12% MeOH, 0.95mM 18C6 5 mM imidazole, 6.75 mM HIBA, pH 4 5 mM pyridine, 3.6 mM tartaric acid, 2 mM 18C6, pH 4 10 mM methylbenzylamine, 15 mM lactate, pH 4.3 5 mM benzimidazole, tartaric acid, pH 5.2, 0.1%HEC,40mM18C6 10 mM p-aminopyridine, acetic acid, pH 4.5, 5 mM 18C6 or 10 mM HIBA 8 mM 4-methylbenzylamine, 15 mM lactate, 5% methanol, pH 4.25

5 mM phthalate, 0.25 mM CTAB, pH 7

8.7 mM benzene-1,2,4-trocarboxylate, pH 4.9

5 mM ATP, 0.02 mM CTAB, pH 3.6 5 mM phthalate, 0.5 mM DTAB, pH 4.2

10 mM phenylphosphonic acid, 200 mM borate, pH 6

5 mM AMP, 100 mM boric acid, 10% water, 80% methanol, 10% acetonitrile

50 mM benzoate to pH 6.2 with L-His, coated capillary

6 mM imidazole, 4 mM formic acid

80 mM Tris, 10 mM benzoic acid

5 mM dihydroxybenzoic acid, 5% methanol, pH 8.1

12 mM 5,5-diethylbarbituric acid, pH 8.6

151 152 153 154

155 156

157 158

159

160

161 162 159 163 164

165

161

166

167

168 169

170

171

172

173

174

175

176


C6SO3 CI EOF

++++++++^•++•l"»•+++•••++++++^.^.++++^'+++•|.+++++-»•+++•f+++++++++++•»•

EOF" +++++++++++++-i-+++-i-+++++++++++++++-i-++++++++++++++++++'f+++

FIGURE 3.18 Illustration of the direction of migration of small ions for a bare silica capillary and a charge-reversed capillary Top and middle: bare silica; bottom: charge-reversed capillary

2. Indirect Detection

The principle of indirect detection is illustrated in Figure 3.19. A UV-absorbing reagent of the same charge (a co-ion) as the solutes serves as an additive to the BGE. This reagent elevates the baseline. When solute ions are present, they displace the additive as required by principle of electroneutrahty As the separated ions migrate past the detector window, they are measured as negative peaks relative to the high baseline.

The criterion for selection of the indirect reagent is as follows:

1. The molar absorptivity of the reagent should be maximized. This means selecting a reagent with a high molar absorptivity and monitoring at the wavelength of maximum absorbance. Selection of a wavelength where the solute does not absorb is equally important. If for some reason a reagent with a modest molar absorptivity must be used, then extended path length capillaries may prove useful (176).

2. The reagent must be ionized at the appropriate pH. Both the reagent and solute must be ionized in order for displacement to occur.

3. The mobility of the reagent should match that of the solutes as closely as possible. Since the reagent concentration must be kept low for sensi-

JNDIRECT ABSORPTION

DARK BUFFER ZONE BRIGHT SAMPLE ZONE DARK BUFFER ZONE

ABS

\

\

\ \ \

V

/ / / /

/ / /

/

TIME

FIGURE 3.19 The principle of indirect absorption detection.

tivity, the problem of electrodispersion is likely to appear. This problem is especially severe since small ions with large diffusion coefficients are being separated. The diagram given in Figure 3.20 permits intelligent

|S203^-

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i . . , L' ,„„._, J,

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i|propkmate''|

.. 1 : 1 etham sulfonate* ] |bmanestitfo.nate'J

1 [propane sulfonati;' |

1 1 S

1 pentane soifonate'

i 1 EELATIVE

i

Cr042-(5.5.103)

i t

1 pyrom«lHtate^-'

1 q.t^io^)

J t

j trimdUtate-^-j (7.1.103)

t 1 phthalate^-(1.4103)

benzoate-(0.8,103)

2000 MIGRATION k kk 'k TIMES

Probes p-phcnolsulfonatc-<0.8.103)

p-toluenesulfonate" (0.3.103)

ip-hydrasybtiijtoatr |(I0.0,103)

FIGURE 3.20 Selection of the indirect reagent based on mobility matching. The relative migration times (chromate = 1) of indirect reagents and solutes (anions) at pH 8 are shown. The numbers in brackets are the molar absorptivities at 254 nm. Reprinted with permission from Trends Anal. Chem., 13, 313 (1994), copyright © 1994 Elsevier Science Publishers.


Table 3.7 Selection of the Indirect Reagent

Anionic Reagents

Chromate^

Phthalate*'

p-Hydroxybenzoate'^

Naphthalene sulfonate^

Pyromellatic acid"

Cationic Reagents

Creatinine

Imidazole

Ephedrine

I -Naphthylamine

Molar Absorbtivity

-

13,000

--

7,800

Molar Absorbti

9,200

5,000^

7,000^

50,000^

ivity

Mobility

High

Medium

Low

Low

High

Mobility (cm^/kV- s)

.38

.44

.25

.12

PK

-

---

5.6^

PK,

-6.9

9.9

3.9

Reference

41

13,41

41

42

11

Reference

19

26

26

26

^Use for common inorganic ions, e.g., chloride. ^Use for carboxylic acids. ' Use for alkyl sulfates or sulfonates. ' At low pH, this material serves as a low-mobility reagent. "Highest ipK^. At 214 nm.

selection of the indirect reagent (177). For example, the highly mobile ions such as chloride, sulfate, nitrate, and mi trite are best detected with either high-mobility chromate or pyromellitate. Organic acids are best detected with either phthalate or benzoate, whereas alkyl sulfonates are best detected with p-toluenesulfonate or even low-mobility naphthalene sulfonate (178). The figures of merit for some indirect reagents are included in Table 3.7.

Normally, no co-ions other than the indirect reagent are present in the BGE; otherwise, sensitivity may be compromised, since displacement of the nonab-sorbing buffer competes with displacement of the indirect reagent. In the separation of alkyl phosphonic acids, 200 mM boric acid is included in the BGE along with the indirect reagent, 10 mM phenylphosphonate. Boric acid is neutral at that pH, and so it does not interfere with detection. The reagent is likely used for secondary equilibrium via complexation with the —OH groups attached to the phosphorus.

The sensitivity of indirect detection is given by (179)


CLOD = ^ , (3.7) (DR)(TR)

where the CLOD is the concentration hmit of detection, CR is the concentration of the reagent, DR is the dynamic reserve, and TR is the transfer ratio. Thus, the lowest CLOD occurs when the reagent concentration is minimal. Ma and Zhang (180) calculated a theoretical CLOD of 2 x 10~^ M for indirect detection using the following equation:

CLOD = ABGE£BGE'^(DR)(TR), (3.8)

Here b is the capillary path length (5 x 10"^ cm), ABGE is the noise of the system (5 X 10-4 AU), £BGE is the molar absorptivity of the reagent (10,000 IVmolcm), the dynamic reserve is 10,000, and the transfer ratio is 1. It becomes possible for the limit of detection of indirect detection to exceed direct detection, at least theoretically, under optimal conditions.

The dynamic reserve is the ratio of the absorbance of the reagent to the noise of the system. The transfer ratio is related to the ratio of the charge on the solute relative to the reagent. For example, chloride should displace a single benzoate ion, but a transfer ratio of only 0.184 has been reported (177). Benzoate is not a good choice for measuring chloride. The best or most sensitive indirect reagents are indicated by the product of the transfer ratio and the molar absorptivity (177). In this regard, chromate, pyromellitate, and trimellitate are the best choices for small anions.

Reagent kits for anions and cations are available from Waters and Hewlett-Packard. These kits are convenient, particularly for scientists new to these techniques. Manufacturers' protocols and support are available with these kits.

3. Secondary Equilibrium

The ionic equivalent conductance (lEC) is useful for predicting separation, since this parameter correlates with mobility (181, 182). As seen in Figure 3.21, ions such as lithium, sodium, and potassium are easily separated. Other metal ions, including transition metals and lanthanides, may have similar lECs, so that com-plexing reagents are required. Since indirect detection is required, the complex-ing reagent must be anionic to avoid interference. Reagents such as a-hydroxyisobutyric acid (HIBA) (181), citrate (181), lactate (183), glycolate (162), and tartaric acid (159) have been used.

Lin et a\. (184), reported on 10 different complexing reagents and found the optimal pH was equal to the ^K^ of the complexing acid. Lactate, succinate, hydroxyisobutyrate, and malonate gave the best performance. Shi and Fritz (161) separated 27 metal cations using 15 mM lactate and 8 mM 4-methylben2ylamine,


Equivalent Ionic Conductance

3 4 4 5 6 7 8 9 10 11

Metals Placed in Ascending EIC

FIGURE 3.21 Plot of the equivalent ionic conductance for the alkah metals, alkaU earth metals, transition metals, and lanthanides. Data from reference (181).

5% methanol in 6 min. Crown ethers can be employed as well in cation separations (185-189). Resolution of potassium/ammonia and calcium/strontium is readily accomplished with 18-crown-6. Methanol can also be used to lower the EOF to improve resolution (183). Lowering the pH for cation separations is also effective in lowering the EOF to improve resolution. A number of these features are employed in the separation of some metal ions, as shown in Figure 3.22. In this case, Cu " is used as the indirect reagent. The mobility of Cu ^ better matches the mobility of the later eluters such as magnesium, strontium, lithium, and barium, as evidenced by the symmetrical peaks.

Most anions have sufficiently differing lECs so that separations are possible without special additives. The EOF, along with the resolution, can be adjusted by altering the concentration and type of added cationic surfactant. Increasing the indirect reagent concentration tends to improve resolution at the expense of sensitivity

A typical anion standard mixture is shown in Figure 3.23. In this case, the indirect reagent is chromate. The mobility of chromate better matches the mobility of the more mobile anions, as shown by the symmetrical peaks. Elec-trodispersion is a normal effect in electrophoresis. Quantitative results are still maintained as long as peak areas are used and sufficient resolution is designed into the separation.


2 3 Minutes

FIGURE 3.22 Separation of cations using Cu " as the indirect reagent. Capillary: 50 cm x 50 lam i.d.; BGE: 4.0 mM copper sulfate, 4.0 mM formic acid, 4.0 mM 18-crown-6; injection: gravity, 10 cm for 10 s; voltage: 20 kV; detection: indirect UV, 215 nm. Key: (1) ammonium, 3.6 ppm; (2) potassium, 7.8 ppm; (3) sodium, 4.6 ppm; (4) calcium, 4.0 ppm; (5) magnesium, 2.4 ppm; (6) strontium, 15 ppm; (7) lithium, 0.69 ppm; (8) barium, 27 ppm. Courtesy of Dionix Corporation.

4. Stacking

Stacking will be covered in detail in Section 8.6. To optimize sensitivity, stacking coupled with electrokinetic injection may be required. The advantages and pitfalls of electrokinetic injection will be covered in Section 8.3. In Figure 3.24, sub-ppm limits of detection are obtained by the use of a 45-s electrokinetic injection at 5 kV This technique works only when the ionic strength of the sample is very low

An advanced technique called transient isotachophoresis, covered in Section 8.6, can be used to further lower the limit of detection (190). In this case, 7 mM chromate, 0.5 mM TTAB, and 1 mM monosodium carbonate was employed as the BGE. The difference in this protocol is the addition of octane sulfonate to the sample to serve as the terminating electrolyte. Under the correct conditions

no Chapter 3 Capillary Zone Electrophoresis

mV

3.0

2.8-j

2.6-]

2.4 H

2.2 i

2.0

1.8

1.6 j

1.4-1

1.2

1.0-1

0.8-j

0.6 i

3.00 3.50 4.00 4.50 Minutes

5.00 5.50

FIGURE 3.23 Separation of anions using chromate as the indirect reagent. Capillary: 60 cm X 75 |Lim i.d.; BGE: 4 mM chromate with 0.3 mM OFM Anion BT; voltage: -15 kV; injection: gravity, 10 cm for 30 s; detection: indirect UV, 254 nm. Key: (1) bromide, 4 ppm; (2) chloride, 2 ppm; (3) sulfate, 4 ppm; (4) nitrite, 4 ppm; (5) nitrate, 5 ppm; (6) fluoride, 1 ppm; (7) phosphate, 4 ppm. Courtesy of Waters Chromatography.

and when the sample is sandwiched between a leading electrolyte and a terminator, a 500-1000 fold trace enrichment can occur. An extended path length capillary (bubble factor 3)^ was shown useful here as well.

5. Direct Detection of Small Ions

A number of ions absorb in the UV region and thus are appropriate for direct injection (129, 191-196). Because of their importance in biological and environmental samples, nitrate and nitrite are the subject of most of these references. Phosphate buffer, pH 3 with detection at 214 nm is a typical operating condition. A coated capillary or charge-reversing cationic surfactant is usually necessary for high-speed separations. An MECC (Chapter 4) separation of bromide, bromate, iodide, iodate, nitrate, and nitrite using DTAB as the surfactant is noteworthy (197). Using an extended path length capillary, a ninefold improvement

^Hewlett-Packard. See Section 9.6 for details.

3.6 Applications and Techniques

-1.0-

111

mAU

3 4 i W^MS/V«*^

2 Minutes

FIGURE 3.24 Separation of trace anions in power plant boiler water using a stacking electroki-netic injection. Capillary: 50 cm x 50 /im i.d.; BGE: 2.25 mM pyromellatic acid, 6.5 mM NaOH, 1.6 mM triethanolamine, 0.75 mM hexamethonium hydroxide; injection: 5 kV for 45 s; voltage: 30 kV; detection: indirect UV, 250 nm. Key: (1) chloride, 15.8 ppb; (2) sulfate, 11 ppb; (3) nitrite, 1.7 ppb; (4) azide; (5) fluoride, 13.3 ppb; (6) formate. Courtesy of Dionix Corporation.

in sensitivity was observed. Molybdate(VI), vanadate(V), and chromate(VI) could be detected at 254 nm in a 0.15 mM CTAB charge-reversed capillary using 6 mM phosphate, 2 mM citrate with detection at 254 nm (198).

Metal complexes formed either precapillary or on-capillary can be detected directly (199-204). Separations are by CZE or MECC. Typical reagents include 4-(pyridylazo)resorcinol (PAR) (205), and CLODs of 10-^ to lO'^M have been reported. Other reagents such as 8-hydroxyquinoline-5-sulfonic acid (206), 1,2-cyclohexanediamine-N,N,N',N'-tetraacetic acid (CyDTA) (207), and EDTA (208) have also been reported.

6. Indirect Fluorescence Detection

Indirect fluorescence detection (209-212) is less frequently used than indirect absorption detection, since few commercial instruments have that capability High-sensitivity analyses are possible in this mode, because it is possible to reduce the concentration of the indirect reagent to very low levels and thus decrease the CLOD predicted by Eq. (3.7). Using 100-|im fluorescein, a mass limit of detection of 20 aM was reported for lactate and pyruvate in single red blood cells (212). Fluorescein is a good additive because is absorbs at 488 nm, the emission wavelength of the argon-ion laser. Electrodispersion is unimportant here since the


solute concentration is so low. At higher solute concentrations, the system will be less useful because of electrodispersion. Of course, the concentration of the indirect reagent could be increased, but then indirect absorption detection becomes applicable.

7. Conductivity Detection

The first reports of conductivity detection for HPCE appeared in the literature in 1987 (213). Seven years later, the first commercial conductivity detector became available.^ A capillary with a fiber-optic type connector and an outlet capillary with the detection sensors are connected to the detection cell assembly. A narrow 24-|Lim spacing is maintained between the two capillaries to minimize band broadening and provide consistent capillary-to-capillary results. This design isolates the cell from the high-voltage circuit (214).

A conductivity detector measures the differences in conductivity between the BGE and each solute. Since high-mobility ions have high conductivity, the use of a low-conductivity BGE provides for sensitive measurements. A typical low-conductivity electrolyte for anions is 50 mM CHES, 20 mM lithium hydroxide, and 0.03% Triton X-100. The charge on the capillary wall is reversed by flushing the capillary with 1 mM CTAB prior to each analysis in order to reverse the EOE The CTAB is often left absent from the BGE to minimize its conductivity. For cations, the BGE is 30 mM histidine, 30 mM MES, and 1 mM 18-crown-6.

The limits of detection for the early eluting, high-mobility solutes are 10 times lower than for indirect detection. For later eluting ions the advantages begin to disappear. This is illustrated in Figure 3.25, where the later eluting ions have a lower response despite solute concentrations that are 5 to 10 times as high as the high-mobility ions. For high-mobility ions, in conjunction with tlTP, sub-ppb limits of detection have been reported (215).

For the determination of low-mobility ions such as alkyl sulfonates or quaternary amines, indirect conductivity is employed to maximize sensitivity (216). In this example, a high-conductivity electrolyte containing 30 mM sodium fluoride and 1 mM triethanolamine is used. Though both cationic and anionic surfactants could be measured in a single run, the LODs were not as good as with indirect detection. To improve the LOD, a higher conductivity BGE would be required, but heating problems would quickly ensue.

B. PEPTIDE MAPPING

Tryptic digests are often employed to determine if changes to a complex protein, such as posttranslational modifications, have occurred. There are many

^Crystal 1000 CE conductivity detector, ATI Unicam (now Thermoquest), Santa Fe, NM.


7.0 Minutes

9.0 11.0

FIGURE 3.25 Separation of anions with conductivity detection. Capillary: 60 cm x 50 |Lim i.d. (ConCap, ThermoQuest); BGE: 50 mM Ches, 20 mM lithium hydroxide, 0.03% Triton X-100; injection: 300 mbs. The capillary is preconditioned with 1 mM CTAB prior to each run. Reprinted with permission from Amer Lah., 28, 25 (1996), copyright © 1996 Int. Sci. Commun.

examples using this technique in the hterature. A series of electrolyte recipes is given in Table 3.8.

Capillary zone electrophoresis is complementary to HPLC for peptide mapping studies (21, 223). Microheterogeneity not detected by HPLC can be resolved by CZE, even for glycoprotein fragments (224). Tryptic digests are usually run by gradient elution reverse-phase liquid chromatography (Figure 3.26a). Run times of several hours are commonplace. By CZE, a run can be completed in 12 min (Figure 3.26b) with modest resolution (225). Lengthening the run time to 60 min further improves the resolution (data not shown). Speed is again the compelling advantage. This feature is conducive to screening large numbers of samples searching for variants, decomposition products, or posttranslational modifications. The CZE mechanism of separation bears no relationship to that of reversed-phase LC. A scatter plot comparing LC retention time with CZE migration time (not shown) for the individual peaks in Figures 3.26a and b yields a random distribution.

CZE has some significant disadvantages compared with HPLC for tryptic mapping:

1. The overall peak capacity of CZE can be lower than gradient elution LC. 2. The difficulty of fraction collection (Section 9.10) means that peak iden

tification through protein sequencing can be a problem. 3. The reproducibility of CZE is not as robust as gradient elution LC.

1 1 4 Chapter 3 Capillary Zone Electrophoresis

Table 3.8 Buffer Recipes for Peptide Mapping

Analyte

Anti-Rh(D) monoclonal antibody

Cytochrome c

Erythropoietin

Globin, from hemoglobin

Human growth hormone

j3-Lactoglobulin

Peptides

Porcine pro-growth hormone releasing hormone

Electrolyte

30 mM phosphate, pH 2.5

25 mM citrate, pH 4

40 mM phosphate, pH 2.5, 100 mM heptanesulfonic acid


100 mM tricine, 30 mM morpholine

5% formic acid, 0.02% Tween 20

100 mM hexane sulfonate, 30 mM phosphate, pH 2.5


Reference

217

218

219

220

21

221

148

222

Nielsen and Rickard (21) reported an empirical method for optimizing a complex separation of human growth hormone (hGH) tryptic digest fragments. This complex sample contains fragments ranging in size from 1 to 32 amino acid residues. Fragments 6-16 and 20-21 consist of two chains connected by a disulfide bond. Fragments 1 and 3 are basic peptides likely to exhibit wall interactions. Both hydrophobic peptides (fragments 4, 6-16, 9, and 10) and hydrophilic peptides (fragments 3, 5, 7, 14, and 17) are present in the sample.

First, it is necessary to ensure adequate buffer ionic strength. Separations in 10 and 100 mM Tricine, pH 8.1, are shown in Figure 3.27. Since pH 8.1 is close to the pi of Tricine, the higher buffer concentration did not draw a very high current. The impact of buffer concentration is substantial due to loading effects, reduction of wall effects, and reduction of the EOF Adding sodium chloride to further increase the ionic strength was not useful.

Four different buffer pH values (2.4, 6.1, 8.1, and 10.4) were initially studied. The best separations were found at pH 2.4 (data not shown) and 8.1, though in both cases a number of fragments were found to overlap. The separation at pH 8.1 was 2.5 as time fast as that at pH 2.4, and so the higher pH was considered a better choice.

Addition of an amine modifier—in this case, morpholine—further improved the resolution (Figure 3.28, p.118), presumably by reducing wall interactions between the peptide fragments and free silanol groups.

Before a separation can be fine-tuned, freedom from wall effects and sufficient buffer capacity are prerequisites. It makes little sense to proceed without these features accounted for. The insufficient resolution at the beginning of the separation calls for more experimental work. Perhaps with the addition of


9S2.659

0.000 SO' 100

Time (mln)

FIGURE 3.26A Reversed-phase HPLC of peptide fragments of a tryptic digest of rhGH. Tryptic fragments are numbered sequentially from the N terminus of the protein and the suffix "c" indicates a peptide fragment resulting from a chymotrypsin-like cleavage. Column: 150 x 4.6 mm i.d. Nucleosil Cis; temperature: 35°C; Mobile phase: A = 0.1% TFA in water; B = 100% acetonitrile; gradient: 100% A, hold 5 min, linear ramp to 37% B over 120 min, linear ramp to 57% B over 10 min; flow rate: 1 mlVmin; detection: UV, 214 nm; injection size: 200 /iL. Reprinted with permission from J. Chromatogr., 480, 379 (1989), copyright ©1989 Elsevier Science Publishers.

cyclodextrins, surfactants, metal ions, or sulfonic acids, complete resolution in a single run may possible.

It should be noted that no single set of conditions resolved all of the tryptic fragments. The same was true for gradient elution LC, even with a 2-h gradient run. At pH 2.4, overlapping fragments 1, 13, 14, and 17 are all resolved.


M I -

9.172

6' "S' 10^ Time (min)

12

FIGURE 3.26B CZE of peptide fragments of a tryptic digest of rhGH. Capillary: polyacrylamide-coated fused silica, 20 cm x 25 im i.d.; buffer: phosphate buffer, pH 2.5; injection: 5 s at 8 kV; detection: UV 200 nm. Peaks were identified by injection individual fractions from the HPLC separation. Reprinted with permission from J. Chromatogr., 480, 379 (1989), copyright © 1989 Elsevier Science Publishers.

C. HIGH-PERCENTAGE ORGANIC AND

NONAQUEOUS ELECTROPHORESIS

Electrophoresis is a process primarily designed for the separation of water-soluble biomolecules. The aqueous systems used in HPCE are advantageous particularly with regard to toxicity and safety Water is not the only solvent that can be used in electrophoresis. Nonaqueous systems may be considered when

I <

Mj 100 150 200 250 300 350 400 4S0 500 550 600

Time (s)

300 350 400 450 500 550 Time (s)

600 650 700 750 800

FIGURE 3.27 CZE of hGH tryptic digest in pH 8.1 tricine buffer. (A) 10 mM and (B) 100 mM. Capillary: 100 cm (80 cm to detector) X 50 jXm i.d.; voltage: 30 kV; temperature: 30°C; detection: UV, 200 nm; injection: 3 s vacuum (10 nL); sample concentration: 90 mM for each fragment (2 mg/mL). Reprinted with permission from J. Chromatogr., 516, 99 (1990), copyright © 1990 Elsevier Science Publishers.

1. The solute is insoluble in water or surfactant-containing solutions, or it forms aggregates.

2. Mass spectroscopy is used, since surfactant-containing electrolytes can be avoided and advantage can be taken of the more efficient nebuliza-tion of organic solvents due to their low surface tension and volatility.

8

o

<

— I —

725 — I —

775

— I —

825 — I

875 375 425 475 525 575 625 Time (s)

— I —

675

FIGURE 3.28 CZE of hGH tryptic digest in pH 8.1 Tricine with 30 mM morpholine. Other conditions as per Figure 3.27. The peaks marked with a * are degradation products. Reprinted with permission from J. Chromatogr, 516, 99 (1990), copyright © 1990 Elsevier Science Publishers.

3. Micropreparative separations are performed, since the low conductivity of the organic solvent permits wide-bore capillaries to be used without experiencing heating problems. For example, a BGE consisting of ethanol: acetonitrile: acetic acid (49:50:1) with 20 mM ammonium acetate allows 200-|lm-i.d. capillaries to be used (226). The loading capacity is 16 times as great as with a 50-|Lim-i.d. capillary.

The characteristics of a good nonaqueous solvent include the ability to dissolve salts (for some conductivity), water solubility (since samples may contain some water), and little absorbance in the UV region of the spectrum (so detection is not limited). While solvents such as formamide may be optimal with regard to electrophoresis, problems with UV absorption and hydrolysis (227) preclude that material from general use. The physical properties of some potential solvents are given in Table 3.9.

An expression for mobility is

Mep 2£C

377 (3.9)

where e is the dielectric constant, ^ is the zeta potential, and rj is the viscosity. For a given ion, the highest speed separations can occur when e/^ is greatest (228). This points to solvents such as N-methylformamide and acetonitrile.


Table 3,9 Characteristics of Solvents for Nonaqueous Electrophoresis

Solvent

Water

Formamide

N-Methylformamide

N,N-Dimethylacetamide

Acetonitrile

Acetic acid

Methanol

bp (°C)

100

210

182

166

82

118

65

Viscosity (cp)

0.89

3.3

1.65

0.78

0.34

1.1

0.54

Polarity

10.2

9.6

6.0

6.5

5.8

6.0

5.1

Dielectric

80

111

182

37.8

37.5

6.2

32.7

£lr\

89.9

33.6

110.3

48.5

110.3

60.6

Data are from reference (227), except e/r\ values, which are from (228).

Since acetonitrile does not absorb in the low UV, it is a solvent worth trying when nonaqueous solvents are called for. N-Methylformamide also has a high e/C, ratio, but the solvent absorbs strongly in the low UV.

Table 3.10 lists some applications that use either a high percentage of organic solvents or totally nonaqueous systems. It becomes easy to question why organic solvents are needed for some of these separations, in particular small inorganic ions such as chloride, nitrate, or fluoride. However, it may become necessary to measure the species in materials such as fuels, solvents, and lubricating oils (229). It has also been reported that the selectivity of separations may be altered by nonaqueous solvents, but the effects cannot be predicted in advance (230).

D. CARBOHYDRATES

Carbohydrate separations fall into several broad categories, including simple sugars, oligosaccharides, and polysaccharides. Glycoproteins may be separated as intact molecules, or the sugars may be released prior to separation. Complex carbohydrates such as glycosoaminoglycans (GAGs) are usually broken down with enzymes to disaccharides prior to separation. The analytical problems to be solved involve both separation and detection. Carbohydrates absorb poorly in the UV, and so indirect detection and derivatization are important techniques. It is useful to determine if more complex detection schemes are required, since high sensitivity may not be an issue for many applications.

Buffer recipes for a variety of carbohydrate samples and detection techniques are given in Table 3.11. Note that several applications employ MECC as the mode of separation. A primer on carbohydrate separations by HPCE (243), a detailed review (244), and a volume of the journal Electrophoresis devoted this subject (245) have appeared since 1994.

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Simple sugars can be separated in complexing borate electrolytes with direct detection at 195 nm. In the absence of borate, the molar absorptivities are too low to be useful. A 50 mM borate BGE in a bare silica capillary maintained at 60°C provides the best results (145). The limits of detection (LOD) are estimated to be around 0.5 mM, which is not very sensitive. Carbohydrates with an unsaturated uronic acid residue absorb more strongly at 232 nm (244).

In the absence of borate, high pH is required to partially ionize the sugars, since the pK^ of most simple sugars is between 12 and 13. An exception is found for sulfated disaccharides, which can be separated under acidic conditions (244). Other complexation chemistries can be employed to improve detection. For GAGs such as heparin, the addition of 5 mM Cu(II) to the BGE (20 mM phosphate, pH 3.5, 240 nm detection) dramatically improves the detection of the extremely microheterogeneous intact heparin molecules (246). Large polysaccharides such as starches are easily detected at 560 nm as the starch-iodine complex (247).

Indirect detection can yield superior sensitivity compared with direct detection. At very high pH, the predominance of hydroxide obscures indirect detection (248). At pH 12.2, the optimal compromise is found. An LOD of 50 |LlM was reported (158) using 25-|Lim-i.d. capillaries with 12 mM riboflavin as the indirect reagent and 63 mM lithium hydroxide to adjust pH. The narrow-bore capillary was required to overcome the heating effects from the high-pH electrolyte. A separation is shown in Figure 3.29. Indirect fluorescence detection (249, 250) using lasers further lowers the LOD, since the concentration of the indirect reagent can be kept very low.

Derivatization techniques improve detection by the addition of a good chro-mophore or fluorophore. This is particularly important when determining the carbohydrate profiles of glycoproteins. Simultaneously, electrophoresis can be improved as well, since negative charges can accompany the derivatizing reagent. Figure 3.30 compares the separation of a dextran standard with three different derivatizing reagents: 2-aminopyridine, 5-aminonaphthalene-2-sul-fonate, and 8-aminonaphthalene-l,3,6-trisulfonate, otherwise known as ANTS (251). The highly mobile ANTS derivatives are advantageous from the standpoint of short analysis time and reduction or elimination of wall effects. Aminopyridine is neutral at the separation pH, whereas the monosulfonate naphthalene derivative contains only one negative charge. Another reagent, 8-aminopyrene-l,3,6-trisulfonate (APTS) proved simpler to use, since the derivative absorbed at the wavelength of the argon-ion laser. In addition, the molar absorptivity and fluorescence quantum yield of the reagent was superior to that of ANTS (252). A helium-cadmium laser was required for excitation of ANTS, and that laser is not part of a commercial system.

For those with a UV detector, 4-aminobenzoic acid ethyl ester reacts with many sugars (253). Separations are performed with 200 mM boric acid titrated

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FIGURE 3.29 Indirect detection of sugars. Capillary: 78 cm x 25 |im i.d.; BGE: 12 mM riboflavin, 63 mM lithium hydroxide; injection: 10 s (ABI270A); voltage: 10 kV; detection: indirect Uy 267 nm; Key: 1 mM solutions of (1) sucrose; (2) maltose; (3) glucose; (4) fructose. Reprinted with permission from J. ChromatogK, A, 716, 231 (1995), copyright © 1995 Elsevier Science Publishers.

to pH 10.5, with detection at 305 nm. The CLOD is an order of magnitude improved compared with that for indirect detection.

E. SERUM PROTEINS

Serum proteins are traditionally separated in the clinical lab by agarose-gel electrophoresis. The patterns obtained are diagnostic for various disease states. Use of an alkaline, high-ionic-strength buffer produces high-speed separations of important serum proteins in an untreated fused-silica capillary (108, 264). A 20 cm x 25 |im i.d. capillary is employed to reduce the heating effects of the high field strength in conjunction with 150 mM borate buffer,io pH 10. The separations (Figure 3.31, top) are reproducible and are similar to the agarose-gel patterns. The run time can be as short as 90 s. Increasing the buffer's ionic strength lowers the EOF and improves the resolution (Figure 3.31, bottom). Multicenter studies evaluating this buffer system on the commercial clinical analyzer have been reported (265, 266).

i^The exact buffer composition was not disclosed.


-J%MM1^

iJi W ^^w

100 iiV 2S}iV

10 SO flain

FIGURE 330 Influence of the fluorescent tag on the separation of a dextran standard with an average molecular weight of 18,300. The reagents used were (A) 2-aminopyridine, (B) 5-aminonaph-thalene-2-sulfonate, and (C) 8-aminonaphthalene-l,3,6-trisulfonate. Capillary: 35 cm x 50 jxm i.d. polyacrylamide-coated; BGE: 100 mM Tris-borate, pH 8.65; field strength: -500 V/cm; detection: LIF, He-Cd at 325 nm excitation, emission at 375 for A, 475 nm for B, and 514 nm for C. Reprinted with permission from Anal. Chem., 66, 1134 (1994), copyright €> 1994 Am. Chem. Soc.


60 70 time (seconds)

1

1 11 /''' l In /

| 9 1 1

1 ... ,..7^**~"'''^''i8^^

150 2OT tjRic (seconds)

FIGURE 3.31 CZE protein profile of a normal control serum. Capillary: 25 cm X 25 im i.d.; voltage: 20 kV; (top) buffer: (proprietary, pH 10.0, probably 150 mM borate); (bottom) higher ionic strength, pH 10; temperature: 22°C; detection: UV, 200 nm. Key: (1) DMF; (2) /-globulin; (20 complements; (3) transferrin; (4) ^-lipoproteins; (5) haptoglobin; (6) a2-"^^croglobuhn; (7) Ofi-anti-trypsin; (8) ai-lipoproteins; (9) albumin; (10) prealbumin. Reprinted with permission from J. Chromatogr., 559, 445 (1991), copyright © 1991 Elsevier Science Publishers.


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140. Mosher, R. A. The Use of Metal Ion-Supplemented Buffers to Enhance the Resolution of Peptides in Capillary Zone Electrophoresis. Electrophoresis, 1990; 11:765.

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143. Wright, R B., Dorsey, J. G. Silver(I)-Mediated Separations by Capillary Zone Electrophoresis and Micellar Electrokinetic Chromatography: Argentation Electrophoresis. Anal. Chem., 1996; 68:415.

144. Plocek, J., Chmelik, J. Separation of Disaccharides as Their Borate Complexes by Capillary Electrophoresis with Indirect Detection in the Visible Range. Electrophoresis, 1997; 18:1148.

145. Hoffstetter-Kuhn, S., Paulus, A., Gassmann, E., Widmer, H. M. Influence of Borate Com-plexation on the Electrophoretic Behavior of Carbohydrates in Capillary Electrophoresis. Anal. Chem., 1991; 63:1541.

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151. Boden, J., Darius, M., Bachmann, K. Determination of Inorganic and Small Organic Anions in Pure Boric Acid Using Capillary Zone Electrophoresis. J. Chromatogr, A, 1995; 716:311.

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1 3 4 Chapters Capillary Zone Electrophoresis

175. Gibbons, J. M., Hoke, S. H. Capillary Zone Electrophoresis with Indirect UV Detection: Determination of Sodium Dodecyl Sulfate in Simulated Stream Water. J. High Resolut. Chromatogr., 1994; 17:665.

176. Nielen, M. W. E Quantitative Aspects of Indirect UV Detection in Capillary Zone Electrophoresis. J. Chromatogr., 1991; 588:321.

177. Buchberger, W, Cousins, S. M., Haddad, P. R. Optimization of Indirect UV Detection in Capillary Electrophoresis of Low-Molecular Mass Anions. Trends Anal. Chem., 1994; 13:313.

178. Romano, J., Jandik, P., Jones, W. R., Jackson, P E. Optimization of Inorganic Capillary Electrophoresis for the Analysis of Anionic Solutes in Real Samples. J. Chromatogr, 1991; 546:411.

179. Yeung, E. S., Kuhr, W. G. Indirect Detection Methods for Capillary Separations. Anal. Chem., 1991; 63:275.

180. Ma, Y., Zhang, R. Optimization of Indirect Photometric Detection of Anions in High-Perfor-mance Capillary Electrophoresis. J. Chromatogr, 1992; 625:341.

181. Weston, A., Brown, P R., Jandik, P, Jones, W R., Heckenberg, A. L. Factors Affecting the Separation of Inorganic Metal Cations by Capillary Electrophoresis. J. Chromatogr, 1992; 593:289.

182. Jones, W R., Jandik, R Controlled Changes of Selectivity in the Separation of Ions by Capillary Electrophoresis. J. Chromatogr, 1991; 546:445.

183. Shi, Y., Fritz, J. S. New Electrolyte Systems for the Determination of Metal Cations by Capillary Electrophoresis. J. Chromatogr, A, 1994; 671:429.

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1 3 6 chapters Capillary Zone Electrophoresis

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219. Rush, R. S., Derby P L., Strickland, T. W, Rohde, M. E Peptide Mapping and Evaluation of Glycopeptide Microheterogeneity Derived from Endoproteinase Digestion of Erythropoietin by Affinity High-Performance Capillary Electrophoresis. Anal. Chem., 1993; 65:1834.

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221. Chang, H.-T., Yeung, E. S. On-Column Digestion of Protein for Peptide Mapping by Capillary Zone Electrophoresis with Laser-Induced Native Fluorescence. Anal. Chem., 1993; 65:2947.

222. Chang, J. P, Smiley D. L., Coleman, M. R. Free-Solution Capillary Electrophoresis of Tryp-tic Digest Fragments of a Recombinant Porcine Pro-Growth Hormone Releasing Hormone (2-76) OH. J. Liq. Chromatogr, 1994; 17:1899.

223. Nielsen, R. G., Riggin, R. M., Rickard, E. C. Capillary Zone Electrophoresis of Peptide Fragments from Trypsin Digestion of Biosynthetic Human Growth Hormone. J. Chromatogr, 1989; 480:393.

224. Nashabeh, W, Rassi, Z. E. Capillary Zone Electrophoresis of apacid Glycoprotein Fragments from Trypsin and Endoglycosidase Digestions. J. Chromatogr, 1991; 536:31.

225. Frenz, J., Wu, S.-L., Hancock, W. S. Characterization of Human Growth Hormone by Capillary Electrophoresis. J. Chromatogr, 1989; 480:379.

226. Valko, I. E., Porras, S. P, Riekkola, M.-L. Capillary Electrophoresis with Wide-Bore Capillaries and Non-aqueous Media. J. Chromatogr, A, 1998; 813:179.

227. Sahota, R. S., Khaledi, M. G. Nonaqueous Capillary Electrophoresis. Anal. Chem., 1994; 66:1141.

228. Valko, I. E., Siren, H., Riekkola, M.-L. Capillary Electrophoresis in Nonaqueous Media: An Overview. LC/GC, 1997; 15:560.

229. Salimi-Moosavi, H., Cassidy R. M. Capillary Electrophoresis of Inorganic Anions in Nonaqueous Media with Electrochemical and Indirect UV Detection. Anal. Chem., 1995; 67:1067.

230. Chiari, M., Kenndler, E. Capillary Zone Electrophoresis in Organic Solvents: Separation of Anions in Methanolic Buffer Solutions. J. Chromatogr, A, 1995; 716:303.

231. Lu, W, Yang, G., Cole, R. B. Determination of Amino Acids by Online Capillary Elec-trophoresis-Electrospray Ionization Mass Spectrometry. Electrophoresis, 1995; 16:487.

232. Cherkaoui, S., Varesio, E., Christen, P, Veuthey, J.-L. Selectivity Manipulation Using Nonaqueous Capillary Electrophoresis. Application to Tropane Alkaloids and Amphetamine Derivatives. Electrophoresis, 1998; 19:2900.

233. Leung, G. N. W, Tang, H. P O., Tso, T. S. C , Wan, T. S. M. Separation of Basic Drugs with Non-aqueous Capillary Electrophoresis. J. Chromatogr, A, 1996; 738:141.

234. Zhang, C.-X., von Herren, E, Thormann, W Separation of Hydrophobic, Positively Chargeable Substances by Capillary Electrophoresis. Anal. Chem., 1995; 67:2070.

235. Wang, E, Khaledi, M. G. Chiral Separations by Nonaqueous Capillary Electrophoresis. Anal. Chem., 1996; 68:3460.

236. Nie, S., Dadoo, R., Zare, R. N. Ultrasensitive Fluorescence Detection of Polycyclic Aromatic Hydrocarbons in Capillary Electrophoresis. Anal. Chem., 1993; 65:3571.

237. Bjoernsdottir, I., Hansen, S. H.J. Chromatogr, A, 1995; 711:313. 238. Tomhnson, A. J., Benson, L. M., Gorrod, J. W, Naylor, S. Investigation of the In Vitro Metab

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241. Ng, C. L., Lee, H. K., Li, S. E Y. Separation of Tamoxifen and MetaboUtes by Capillary Electrophoresis with Non-aqueous Buffer System. J. Liq. Chromatogr., 1994; 17:3847.

242. Naess, O., Tilander, T., Pedersen-Bjergaard, S., Rasmussen, K. E. Analysis of Vitamin Formulations by Electrokinetic Chromatography Utilizing Tetradecylammonium Ions as the Pseu-dostationary Phase. Electrophoresis, 1998; 19:2912.

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250. Garner, T. W, Yeung, E. S. Indirect Fluorescence Detection of Sugars Separated by Capillary Zone Electrophoresis with Visible Laser Excitation. J. Chromatogr, 1990; 515:639.

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252. Evangelista, R. A., Liu, M.-S., Chen, F-T. A. Characterization of 9-Aminopyrene-l,4,6-trisul-fonate-Derivatized Sugars by Capillary Electrophoresis with Laser-Induced Fluorescence Detection. Anal. Chem., 1995; 67:2239.

253. Vorndran, A. E., Grill, E., Huber, C , Oefner, P J., Bonn, G. K. Capillary Zone Electrophoresis of Aldoses, Ketoses and Uronic Acids Derivatized with p-Aminobenzoate. Chromatographia, 1992; 34:109.

254. Zhang, Y, Arriaga, E., Diedrich, P, Hindsgaul, O., Dovichi, N. J. Nanomolar Determination of Aminated Sugars by Capillary Electrophoresis. J. Chromatogr, A, 1995; 716:221.

255. Yu, Z., Chen, Y, Xu, G., Marina, M. L. Separation and Determination of Ganghosides Using High Performance Capillary Electrophoresis. J. Liq. Chromatogr Related Technol., 1998; 21:349.

256. Malsch, R., Harenberg, J., Heene, D. L. High-Resolution Capillary Electrophoresis and Poly-acrylamide Gel Electrophoresis of Heparins. J. Chromatogr, A, 1995; 716:259.

257. Honda, S., Iwase, S., Makino, A., Fujiwara, S. Simultaneous Determination of Reducing Monosaccharides by Capillary Zone Electrophoresis as the Borate Complexes of N-2-Pyridylgly-camines. Anal. Biochem., 1989; 176:72.

258. Le, X. C, Zhang, Y, Dovichi, N. J., Compston, C. A., Palcic, M. M., Beever, R. J., Hindsgaul, O. Study of the Enzyme Transformation of Fluorescently Labeled Oligosaccharides in Human Epidermoid Cells Using Capillary Electrophoresis with Laser-Induced Fluorescence Detection./. Chromatogr, A, 1997; 781:515.

259. Le, X., Seaman, C, Zhang, Y, Zhang, J., Dovichi, N. J., Hindsgaul, O., Palcic, M. M. Analysis by Capillary Electrophoresis-Laser Induced Fluorescence Detection of Oligosaccharides Produced from Enzyme Reactions. J. Chromatogr, A, 1995; 716:215.

260. Klockow, A., Amado, R., Widmar, H. M., Paulus, A. Separation of 8-Aminonaphthalene-l,3,6-trisulfonic acid-Labelled Neutral and Sialylayed N-Linked Complex Oligosaccharides by Capillary Electrophoresis. J. Chromatogr, A, 1995; 716:241.


261. Schaeper, J. R, Shamsi, S. A., Danielson, N. D. Separation of Phosphorylated Sugars Using Capillary Electrophoresis with Indirect Photometric Detection. J. Capillary Electrophor., 1996; 3:215.

262. Kakehi, K., Susami, A., Taga, A., Suzuki, S., Honda, S. High-Performance Capillary Electrophoresis of O-Glycosidically Linked Sialic Acid-Containing Oligosacchardies with Low-Wavelength UV Monitoring. J. Chromatogr., A, 1994; 680:209.

263. Mechref, Y., Ostrander, G. K., El Rassi, Z. Capillary Electrophoresis of Carboxylated Carbohydrates. 1. Selective Precolumn Derivatization of Gangliosides with UV Absorbing and Fluorescent Tags. J. Chromatogr., A, 1995; 695:83.

264. Chen, E-T. A., Liu, C.-M., Hsieh, Y.-Z., Sternberg, J. C. Capillary Electrophoresis—a New Clinical Tool. Clin. Chem., 1991; 37:14.

265. Bienvenu, J., Graziani, M. S., Rpin, E A., Bernon, H., Blessum, C, Marchetti, C, Righetti, G., Somenzini, M., Verga, G., Aguzzi, E Multicenter Evaluation of the Paragon CZE 2000 Capillary Zone Electrophoresis for Serum Protein Electrophoresis and Monoclonal Component Typing. Clin. Chem., 1998; 44:599.

266. Bossuty X., Schiettekatte, G., Bogaerts, A., Blanckaert, N. Serum Protein Electrophoresis by CZE 2000 Clinical Capillary Electrophoresis System. Clin. Chem., 1998; 44:749.

CHAPTER 4 Capillary Zone Electrophoresis Secondary Equilibrium, Micelles, Cyclodextrins, and Related Reagents

4.1 Introduction 4.2 Micelles 4.3 Separation Mechanism 4.4 Selecting the Electrolyte System 4.5 Elution Range of MECC 4.6 Alternative Surfactant Systems 4.7 Cy clodextrins 4.8 Applications and Methods Development 4.9 Chiral Recognition 4.10 Affinity Capillary Electrophoresis

References

4.1 INTRODUCTION

Retention in liquid chromatography is based on the distribution of a solute between two discrete phases, the stationary phase and the mobile phase. A separation between two or more solutes can be achieved whenever the equilibrium distribution between the phases is distinct for each component in the mixture. Under this condition, solute will differentially migrate through a chromatographic column. The separation factor, a, for solutes A and B can be expressed as (1)

139


a = — -—^, (4.1) [A],[B],

where [A] = the concentration of A in phase x or y and [B] = the concentration of B in either of the chromatographic phases.

Nowhere in Eq. (4.1), nor in any of the other fundamental expressions for retention in chromatography, is there an absolute stipulation of the velocity of either phase. It is generally assumed that one phase is mobile while the other is stationary. If that assumption of a stationary phase is disregarded, it is easy to imagine a chromatographic separation taking place through the equilibrium distribution of a solute between two phases that are moving at differing velocities. This concept forms the basis for many forms of electro-kinetic separations.

Chromatographic-type processes in HPCE are the most profound applications of secondary equilibrium in CZE. While these are often considered as separate techniques, they are a variant of CZE, since the BGE is uniform throughout the capillary and electrolyte reservoirs.

The semistationary, or slow-moving, phase in electrokinetic chromatography is composed of molecular aggregates or discrete molecules that are dissolved as additives in the BGE. This formulated buffer system contains, on a molecular level, a heterogeneous environment or "pseudophase" that can compete with the bulk aqueous solution in interacting with the solute. The driving forces that control the speeds of the bulk solution and the heterogeneous pseudophase are electroosmotic and/or electrophoretic migration factors.

The creation of the pseudophase can be accomplished with a variety of buffer additives. Surfactants that generate aggregates known as micelles are the most common additives. This form of EKC is known as micellar electro-kinetic (capillary) chromatography (MEKC or MECC). The first reports of this remarkable advance were published in 1984 and 1985 (2, 3). Microemulsions (4-6), liposomes (7), and vesicles (8) represent additional examples of molecular aggregates acting as a pseudophase. Molecules that are not aggregates can also be considered a pseudophase. Cyclodextrins (9) are representative of that class, but dendrimers (10), macrocyclic antibiotics (11), polymeric ion-exchange reagents (12, 13), and polymeric surfactants (14) can be used in a similar fashion.

Electrokinetic chromatographic separations are used primarily for the separation of small molecules, though there have been reports on the separation of proteins (15,16). Unique applications such as chiral recognition can be accomplished directly with micelles (17), cyclodextrins (18, 19), crown ethers (20), or macrocyclic antibiotics (11) or by the MECC separation of diastereomers that were prepared by precapillary derivatization (21). Each of these techniques will be covered in this chapter.

4.2 Micelles 141

4.2 MICELLES

Surfactants are molecules comprising long hydrophobic "tails" and polar "head groups." Above a certain concentration, known as the critical micelle concentration (CMC), surfactant molecules spontaneously organize into roughly spherical to ellipsoidal aggregates known as micelles. This form of molecular organization occurs due to hydrophobic and electrostatic effects and serves to lower the free energy of the system.

In aqueous solution, the surfactant's hydrophobic tail cannot be solvated by water molecules. As the concentration of surfactant is increased in the bulk solution, the molecules begin to find each other with increasing probability. Since the polar head groups are solvated in aqueous solution, the surfactant molecules orient toward each other's tail, forming first a dimer, and later trimers, tetramers, and so forth. These aggregates are known as premicellar assemblies. Finally, at the CMC, the full micellar organization takes shape, a drawing of which is shown in Figure 4.1. The aggregate forms with a hydrophobic core, the result

FIGURE 4.1 Representation of an anionic micelle associated with a solute, naphthalene. The spiked shapes indicate the anionic head group.


of tail-in orientation. Shape and stability of micelles are further determined by electrostatic repulsion of the polar head groups and van der Waals attraction of the lipoid chains. Materials insoluble in water are frequently dissolved through hydrophobic interaction with the surfactant. With polar head groups at the periphery, electrostatic interaction with external solutes can occur provided ionic surfactants are employed. The micellar model provides for four zones (22):

1. A hydrocarbon-like core with a diameter of 10-28 A 2. The Stern layer, which contains the polar head groups and counterions 3. The Gouy-Chapman layer, an electric double layer that is hundreds of

angstroms thick 4. The bulk surrounding water

At the CMC, the bulk properties of the micellar solution are dramatically altered, including surface tension, conductivity, solubilizing power, and the ability to scatter light. Micelles are dynamic entities in equilibrium with the surrounding environment. Surfactant molecules are free to exchange between the micelle and the external media. Solutes dissolved in surfactant solutions are free to exchange within the micelle as well. For example, typical entrance rate constants for arenes dissolved in sodium dodecyl sulfate are 10^ s"i, whereas the exit rates are lO' s"i.

Another class of surfactants forms micelles in nonaqueous solvents. Known as inverted micelles, these aggregates have an aqueous core, with the hydrophobic portion of the surfactant in contact with the bulk organic solvent. There have been no reports to date of the use of inverted micelles in electrokinetic separations.

Micellar solutions play an important role in many phases of analytical and organic chemistry, including catalysis, electrochemistry, spectroscopy, chromatography, and now, capillary electrophoresis. The use of these intriguing solutions for analytical chemistry has been reviewed (23). Of particular relevance to this chapter is the use of micellar mobile phases in liquid chromatography (24). Surfactant solutions above the CMC can serve as mobile-phase modifiers and function in a similar fashion to conventional organic solvents in reversed-phase liquid chromatography. The phase distribution is more complicated because of the presence of a chromatographic stationary phase, the micellar aggregate, and the bulk aqueous solution. The phase equilibria of a solute between these phases is shown in Figure 4.2. In MECC, the absence of the stationary phase simplifies the phase distribution (25).

Sodium dodecyl sulfate (SDS) is the most widely used surfactant for both electrophoresis and chromatography. This surfactant has the proper hydrophilic-lipophilic balance (HLB) for its intended use. In other words, SDS is very water soluble and has a high degree of lipid-solubilizing power. Because of its widespread applicability, the surfactant is available in highly purified form and is very inexpensive. The CMC for SDS is 8 mM, and its aggregation number is 63 (22). While many surfactants can be employed in electrokinetic sep-

4.3 Separation Mechanism 143

FIGURE 4.2 Partition coefficients for a solute in micellar liquid chromatography. K^ = stationary phase-aqueous phase, K^^ = micellar phase-aqueous phase, and Kgm = stationary phase-micel-lar phase partition coefficients.

arations, much of this chapter, as reflected by the scientific hterature, will be devoted to the use of SDS.

The pioneering works of Jorgensen and Lubacs (26) for CZE and Armstrong and Nome (24) for micellar liquid chromatography, both appearing in 1981, provided pieces of a puzzle, the solution of which led to the discovery and development of MECC.

4.3 SEPARATION MECHANISM

A. BASIC CONCEPTS

In untreated fused silica, the EOF, which is directed toward the cathode, is substantial at pH values ranging from mildly acidic through alkaline. On the other hand, SDS micelles are anionic and electrophorese toward the anode. As a result.


the overall micellar velocity is reduced compared with the bulk flow. These concepts are illustrated in Figure 4.3. Electroosmotic flow overcomes the micellar electrophoretic velocity at the aforementioned pH range, resulting in a net micellar flow toward the cathode.

Since a solute may partition into and out of the micellar aggregate, its own migration velocity can be affected as well. When partitioned into the micelle, solute velocity is retarded. When present in the bulk phase or interstitial space between micelles, the solute, if neutral, is simply swept through the capillary by the EOF This too appears in Figure 4.3, where a mixture of naphthalene, anthracene, and pyrene represent prototypical neutral molecules. In that mixture, naphthalene elutes first, since it spends more time in the bulk aqueous phase.

An illustration of a single-component separation is shown in Figure 4.4. The term t^ is analogous to the chromatographic description of the void volume of the column. Similarly, t describes the retention of a solute. The term t^^^^ which describes the velocity of the pseudophase, distinguishes MECC from chromatography Under most separation conditions, all solutes must elute between t^ and t c-

The fundamental equation for fe' accounts for presence of the mobile pseudophase:

¥ = ^od-^R/fmc)

(4.2)

M ep"

EOF lojp)

FIGURE 4.3 Illustration of the micelle being swept toward the cathode by the EOF while coun-termigrating toward the anode. The solutes are partitioning between the micelle and bulk aqueous phase. Separation occurs due to differences in hydrophobic interaction with the micelle.

DETECTOR

-Ld-

WATER SOLUTE

INJECTOR

MICELLE

i 1

0 IR mc TIME

FIGURE 4.4 Representation of the zones separated in a capillary (upper trace) along with the detected electropherogram (lower trace) for a hypothetical mixture of water, solute, and micelle. The broadening of the slowly migrating peaks is a consequence of on-capillary detection (Section 9.1) and diffusion.

As the velocity of the pseudophase approaches zero (a true stationary phase), t c approaches infinity, and Eq. (4.2) reduces to the classical chromatographic expression for ^'.Equation (4.2) implies that as t^^ is approached, the peaks elute at more closely spaced intervals. Terabe et al. (3) recognized this effect is similar to that obtained with concave gradient elution LC for solutes with fe'< 150. The following equation describes the resolution between two solutes by MECC:

a - 1

V ^ y l + k 2 /

1 - tjt^.

l+(tjt^jk[ (4.3)

As in Eq. (4.2), when the micellar velocity approaches zero, the equation reduces to the classical expression for chromatographic resolution.

The optimal value for k' (maximum resolution) is given by

K,. = (t^JO'"- (4.4)

The parameters tg and t^^ must be determined experimentally (Section 4.5).

146

B. ELUTION ORDER


Prediction of the elution order can be straightforward, provided a homologous series of compounds are being separated. MECC has the capability of separating, within a single run, anionic, neutral, and cationic species. Figure 4.5 shows the separation of a series of peptides that differ only by a single amino acid. Peptide 15 has a net charge of - 2 and is strongly repelled from the anionic SDS micelle. As a result, the peptide spends much of its time in the bulk phase, thereby migrating the fastest of the group. Peptide 2 has a charge of -1 and is repelled less strongly, so it spends more time attached to the micelle and exhibits a longer migration time than does peptide 15. Peptides 1 and 7 are neutral and are separated based on hydrophobic effects.^ Since these peptides are not repelled from the micelle, the migration times are lengthened relative to the anionic peptides. Finally, the cationic peptides are last to elute. These peptides exhibit strong electrostatic interaction with the micelle, and as a result, both have lengthy migration times compared with the other peptides.

Note that the migration order of the charged peptides is the reverse of what is found without the use of the surfactant. In the absence of the micelle, the cationic peptides migrate rapidly toward the cathode, since both the EOF and electrophoretic mobility are in the same direction. The anionic peptides coun-

iThe calculation of neutrality is based on the pH of the bulk solution. Since the pH is much lower at the micellar surface (due to an electrical double layer of protons), it is probable that these "neutral" peptides are cationic in the micellar domain.

»»<<>rt>#S#M>»JI

S D S M E C C

PEPTIDE i 15 AFDfONG

2 AFDAING 1 AFAAING

1 7 AFKADNG 4 AFKAING

! 10 AFKIKNG

CHARGE - 2 - 1

0 0

+1 +2

1 I I I J L_ 10 16 18 20 22 24 26 28

TIME (min)

FIGURE 4.5 MECC of cationic, anionic, and neutral peptides. Capillary: 65 cm (45 cm to detector) X 50 i^m i.d.; BGE: 10 mM phosphate, 100 mM SDS, pH 7.0; voltage: 20 kV; injection: vacuum, 2 s; detection: UV, 200 nm. Courtesy of Applied Biosystems, Inc.


termigrate toward the positive electrode but are swept by the EOF toward the negative electrode. As a result of this, the anionic peptides elute last.

For solutes that are not part of a homologous series, prediction of the elu-tion order is a daunting task. Both electrostatic and hydrophobic interaction with micelles are in force. If the solutes are charged, they to will also experience electrophoresis, at least when contained in the bulk solution.

The structures of a series of nonsteroidal anti-inflammatory drugs are shown in Figure 4.6. These compounds are all aromatic and have carboxylic acid groups as well. Otherwise, phenyl, biphenyl, naphthalene, and other moieties form the structural features of these diverse compounds. Separations by CZE and MECC are shown in Figure 4.7 (27). There is no apparent rationale for the comparative order of migration of these compounds by either mode of electrophoresis. With reversed-phase LC, the order of elution is peak numbers 3, 1, 5, 2, 4. If only hydrophobic effects were in operation by MECC, the order of elution by LC would be expected to be comparable to that by MECC. Since the factors that contribute to the solutes' migration velocity by MECC are complex, a theoretical approach toward the prediction of retention requires a model that considers solute-micelle hydrophobic and electrostatic interactions as well as the solute's electrophoretic properties.

HgCO"

CHgCOOH

CH2CH(CH^2 HgCO' CH2COOH

jmamrnAcm

CH3COOH

TCHJffitTK

FIGURE 4.6 Structures of nonsteroidal anti-inflammatory drugs.


yiLL

TIME(MiN.)

FIGURE 4.7A CZE of non-steroidal antiinflamatory drugs. Capillary: 64.5 cm (43.5 cm to detector) x 25|Ltm i.d., BGE: 20mM phosphate, pH 7.0; 25 mM SDS; voltage: 25kV; temperature: 30°C; injection: vacuum, 2 sec; detection, UV 230 nm. Key: 1) sulindac, 100 jXg/mL; 2) indomethacin, 100 }ig/mL; 3) tolmetin, 100 |ig/mL; 4) ibuprofen, 100 )J.g/mL; 5) naproxen, 10 |ig/mL; 6) diflunisal, 50 |Xg/mL. Reprinted with permission from J. Liq. Chromatogr., 14, 952 (1991), copyright ©1991 Marcel Dekker.

4.4 SELECTING THE ELECTROLYTE SYSTEM

A. SURFACTANT CONCENTRATION

A general recipe for an MECC electrolyte includes the surfactant, usually SDS, a buffer to fix the pH, and other additives to adjust k and/or the overall elution range (t^Jt^). The SDS concentration generally ranges from 25 to 150 mM.

4.4 Selecting the Electrolyte System 149

" TrME(MIN.) ^

FIGURE 4.7B MECC of non-steroidal antiinflammatory drugs. BGE: 20mM phosphate, pH 7.0. Other conditions and key as per Fig. 4.7A. Reprinted with permission from J. Liq. Chromatogr., 14, 952 (1991), copyright ©1991 Marcel Dekker.

Higher SDS concentrations usually result in longer solute migration times, since the probability of partitioning into the micelle increases. Since SDS is ionic, the current increases as well.

As Figure 4.8 indicates, substantial selectivity can be designed into the separation, depending on the degree of interaction between the solutes and the micellar assembly (28). That interaction can be hydrophobic or electrostatic. For example, vitamins B and 3 2 ^^^ cationic, thereby forming ion pairs with the anionic micelle. On the other hand, anionic species are repelled from


.•-vitamin Bl

PL-5'-phosphate

vitamin B12 niacin P[\/l-5'-phospiiate vitamin B2 phosphate vitamin B2

-vitamin B6 \ pyridoxyl (PL)

nicotinamide pyridoxylamine (PM)

0.05 0.1 0.15 SDS CONCENTRATION (M)

FIGURE 4.8 Effect of SDS concentration on the retention time of 11 water-soluble vitamins. BGE: 20 mM phosphate-borate, pH 9.0 plus SDS. Capillary: 65 cm (50 cm to detector) x 50 jim i.d.; voltage: 20 kV; temperature: ambient; detection: UV, 210 nm. Redrawn with permission from J. Chromatogr, 465, 331 (1989), copyright © 1989 Elsevier Science Publishers.

anionic micelles. In this case, increasing the surfactant concentration may not affect the migration time unless hydrophobic interactions are significant.

It is possible to calculate the optimal surfactant concentration (25):

[SURF] = k'

+ CMC, P V

(4.5)

Here [SURF] is the optimal surfactant concentration, P^^ is the partition coefficient of the solute between the water phase and the micellar phase, and V is the partial molar volume of the surfactant. Since it is necessary to experimentally determine some of the parameters of this equation, it is seldom used in practice.

4.4 Selecting the Electrolyte System

B. EFFECT OF P H

151

A suitable buffer is chosen, depending on the required pH. Many papers have reported on the use of a phosphate-borate blend. The advantage of this composition is the maintenance of a common ionic environment over a pH range including 6-11. For reasons not entirely clear, the borate-phosphate blend provided better peak symmetry than a borate-acetate blend for water-soluble vitamins (28). Following that paper, the buffer blend became self-perpetuating. In most cases, borate buffer, pH 9.3, or phosphate buffer, pH 7, is sufficient.

The selection of pH may be based on the pK values of the solutes and the requisite selectivity. The ¥ for neutral compounds is pH independent. For bases, ¥ increases as the pH is lowered due to ion pairing with the anionic SDS micelle. For acids, k'decreases as the pH is raised due to ion repulsion with the micelle. Studying the impact of pH on migration time and selectivity is useful for selecting the optimal pH for the separation of charged solutes. Figure 4.9 illustrates a migration time versus pH plot for several vitamins (28). The separation is best at pH 8.5.

15

1 | 1 0

z o I-

o

8

PL-5*-phosphate

vitamin B1

/iiiaan PM-5*"phosphat0

--vitamin B12 -vitamin B6 pyridoxyl (PL) nicotinamide pyridoxylamine (PI

9 pH FIGURE 4.9 Effect of pH on the retention time of 11 water-soluble vitamins. SDS concentration: 50 mM; voltage: 25 kV Other conditions as per Figure 4.8. Redrawn with permission from J. Chro-matogr., 465, 331 (1989), copyright © 1989 Elsevier Science Publishers.


In addition to the impact of pH on charge, the EOF is also affected. As Figure 4.10 illustrates, this has a profound impact on the technique. Since SDS is ionized at all pH values studied, Vgp has a constant and negative velocity. The electroosmotic velocity Vgo is positive and changes as usual as the pH is adjusted. The net migration velocity v^^ of the SDS micelle is a function of both v^^ and Vgp. At pH 5, the net migration velocity of the micelle approaches zero. At this point, we have a stationary phase. When pH is above 5, SDS migrates toward the cathode; when pH is below 5, its direction of migration reverses.

At low pH, it is necessary run the separation in the reversed-polarity mode. The order of migration, too, is reversed from the high-pH run, since hydropho-

FIGURE 4.10 Impact of pH on the EOF (v^o), the electrophoretic velocity of the SDS micelle (Vep), and the net direction of micellar migration (Vmc)- Reprinted with permission from J. Microcolumn Sep., 1, 150 (1989), copyright © 1989 Microseparations, Inc.

4.4 Selecting the Electrolyte System 153

bic compounds spend more time attached to the micelle and elute first. These features are illustrated in Figure 4.11. To speed up the separations, it is advantageous to use a coated capillary to completely suppress the EOF (29, 30). Since the bulk liquid is stationary, only solutes that are anionic or partition into the micelle will be swept past the detector.

(a)

Jn*MIMMM*|W« L L r~T~rT~T~sr-5~?4

tim* Cininut««)

(b) \2

y 3

U I—I 1 — X — r

4

1—X—^ 30

•^-Ifs I S tlmm (mifiut#s)

FIGURE 4.11 Effect of pH on the order of elution of parabens. Capillary: 100 cm (50 cm to detector) X 100 |im i.d.; BGE: 50 mM SDS, 10 mM phosphate, in a, pH 7.0; in b, pH 3.37; voltage: in a, +25 kV; in b, -25 kV; injection: electrokinetic, in a, +5 kV, 5 s; in b, -10 kV, 10 s; detection: 254 nm; Key: (a) (1) methyl; (2) ethyl; (3) propyl; (4) butyl paraben. (b) (1) butyl; (2) propyl; (3) ethyl; (4) methyl paraben. Reprinted with permission from J. High Res. Chromatogr., 12, 635 (1989), copyright © 1989 Dr. Alfred Heuthig Publishers.


4.5 ELUTION RANGE OF MECC

Micellar electrokinetic separations have a limited elution range, which is defined by the terms to and t^^.

A. MEASUREMENT OF t o

Determination of t^ can be accomphshed by measuring the transit time to the detector for a neutral species that has no affinity for the micelle. Methanol, acetone, or formamide is typically selected.

B. MEASUREMENT OF t mc

The calculation of the capacity factor ^'requires the knowledge of t c the micellar migration time. This is determined by employing a probe such as Sudan 111, a water-insoluble dye that is bound to micelles (3). When organic solvents are used as additives, the probe method becomes insufficient, since the dye can partition into the bulk phase. In this example, the determination of tmc becomes difficult. A homologous series of compounds of increasing hydrophobicity has been employed to determine i^^ by an iterative calculation (31).

In this method, a series of dansylated aliphatic amines was employed, including Ci, C6, Cg, and C12, in a buffer system containing 25% methanol and 25 mM SDS. The migration time of dodecylamine only differed from octylamine by less than a minute despite a four-carbon chain length difference, and so it was assumed that dodecylamine was migrating at a rate close to the micellar velocity. This was tested by plotting log fe' versus the carbon number of all solutes except dodecylamine. A fe'for dodecylamine was then extrapolated. The calculated migration time, assumed equal to t^^^ was used to calculate a new set of K'values using Eq. (4.2). This process was repeated until successive iterations showed no substantial differences in t c- This technique proved that dansylated dodecylamine could be used as a t^^ marker with an error of only 0.04%. Because of these difficulties, it is fortunate that accurate measurement of t^c is seldom necessary.

C. INCREASING THE ELUTION RANGE

The peak capacity of MECC is directly proportional to Init^c^to); therefore, increasing the ratio t^c^t^ will increase the number of components that can be resolved in a single run (32). Decreasing the EOF with a treated capillary is one

4.5 Elution Range of MECC 1 5 5

means of improving this ratio (32, 33). When using Cg- or Cis-coated capillaries, hydrophobic sites on the capillary are effectively saturated by SDS, so that wall binding is not a problem. Binding of SDS to the capillary wall is sufficiently strong that the net surface charge on the capillary remains anionic, though the charge density, as evidenced by the reduced EOF, is lower than that for bare silica.

Since the capillary coatings are usually unstable at high pH, it is better to use other means of increasing the elution range. Organic modifiers can also be used to modify the elution range (34, 35). It is far more productive to consider the use of the modifier as in LC—as a means of adjusting the solute's partition coefficient between the chromatographic phases. The selection of the modifier can increase both to and t^^. The use of methanol or other linear alcohols reduces the EOF, whereas acetonitrile has a much lesser effect.

The full impact of the use of the organic modifier is illustrated in Figure 4.12 (36). These separations are for a series of impurities found in heroin seizure samples. Many of these impurities are very hydrophobic and elute near t^^. The addition of 15% acetonitrile alters the partition coefficients and dramatically lowers fe'for many of these components.

Many organic modifiers are useful in MECC. These include methanol, propanol, acetonitrile, tetrahydrofuran, and dimethylformamide. Acetonitrile has the particular advantage of not affecting the EOF, and the solvent does not absorb in the low UV.

The percent modifier that can be added is limited by the impact of the solvent on the micellar aggregate. Features such as the CMC, aggregation number, and micellar ionization (rate of exchange of surfactant between micelle and bulk solution) are affected by the percent organic modifier. Generally, the use of less than 25% organic modifier does not totally disrupt the micellar aggregate. Higher amounts of modifier may cause sufficient micellar disorder that the separation mechanism is changed. Separations may still occur, due to hydrophobic binding between the nonmicellized surfactant and a solute (37). This technique has been used to separate very hydrophobic compounds such as C20 aryl ketones.

Highly concentrated solutions of urea are frequently employed to solubilize proteins, DNA, hydrocarbons, and amino acids. The mechanism of solubilization is probably due to a diminished water structure surrounding the hydrophobic solute (38). Both to and t c are affected, but the elution window increases, as indicated by the decreasing tjt^^ ratio seen in Table 4.1. In addition, the In V values for hydrophobic solutes such as naphthalene, phenanthrene, and fluoranthene show a linear decrease as the concentration of added urea is increased. The current decreases as well, due to an increase in the viscosity (1.66 times as viscous for 8 M urea) of the electrolyte as well as other changes in ionic mobiUties.

Separations of 23 PTH-amino acids and eight corticosteroids were reported using this technique without the need for organic solvent modifiers (38). Using

ttl O

s m <

(a)

J - A ^ L

T 1 1 1 1 T-16 32 4 8

TIME (min.)

a tso-j

a nan

a 0 3 7 - |

"T I I 1 1 1 1 1 1 1 1 1 O S 1 0 1 S 2 0 2 S 3 0 3 S 4 0 4 5 9 0 S 5

MIKUTES

FIGURE 4.12 Impact of the organic modifier on the MECC separation of heroin impurities. Capillary: in a, 100 cm; in b, 50 cm x 50 |Im i.d.; BGE: (a) 100 mM SDS, 10 mM phosphate-borate, pH 8.5; (b) 85 mM SDS, 8.5 mM phosphate-borate, pH 8.5, 15% acetonitrile; temperature: 50°C; detection: Uy 210 nm. Reprinted with permission from Anal. Chem., 63, 823 (1991), copyright © 1991 Am. Chem. Soc.

4.6 Alternative Surfactant Systems 157

Table 4.1 Migration Times of the Aqueous Phase and Micelle at Different Urea Concentrations

Migration Time Urea Concentration (M)

0.0 2.0 4.0 6.0 8.0

to (min)

t^e (min)

'-c/'-mc

3.92

14.57

0.269

3.92

16.10

0.243

4.65

22.76

0.204

5.46

30.11

0.181

6.38

6.45

0.175

Buffer: 50 mM SDS in 100 mM borate-50 mM phosphate; voltage: 20 kV. Data from reference (38).

computer-aided experimental design, optimization of urea and SDS was accomplished for pesticides, derivatized amines, and nitrotoluenes (39).

Urea-based electrolytes are very useful in keeping marginally soluble materials in solution in a totally aqueous media. Its main problem is absorption in the low-UV region of the spectrum. Since urea absorbs strongly below 230 nm, the solutes must absorb light above that wavelength.

D. DECREASING THE ELUTION RANGE

Admixtures of SDS and a nonionic surfactant such as Brij-35 (polyoxyeth-ylene-23-lauryl ether) can be employed to form co-micelles in solution. Since Brij-35 is neutral, the mobility of the co-micelle is lower than that of the pure SDS micelle. Thus, t^^ is decreased, while t^ is unchanged. Concomitant with the reduction in the elution range is a change in selectivity (40-42). While the effects on selectivity cannot be predicted in advance, it is very worthwhile to try this approach. Typical electrolytes for this combination are 20 mM borate, pH 9.3, 25-100 mM SDS, and 10-50 mM Brij-35. The efficiency of the separation can be 2-3 times as great as that with SDS alone (40). Optimization of the system is simple once scouting runs give an indication of separation.

4.6 ALTERNATIVE SURFACTANT SYSTEMS

The number of potential reagents for MECC is enormous and overwhelming. The reader should be aware that these alternative surfactant systems represent less than 10% of the world's literature on MECC. A partial listing of surfactants is given in Table 4.2.


Table 4.2 Surfactants for MECC

Anionic CMC(mM)

Sodium dodecyl sulfate (SDS) 8

Sodium decyl sulfate (STS) 40

Sodium taurocholate (STC) 10-15

Sodium cholate (SC) 13-15

Sodium taurodeoxycholate (STDC) 2-6

Sodium deoxycholate 4-6

Sodium lauroyl methyltaurate (SLMT)

Catonic

Decyltrimethylammonium chloride (DTAC) or bromide (DTAB) 61, 68

Dodecyltrimethylammonium chloride (DoTAC) or bromide (DoTAB) 20. 16

Cetyltrimethylammonium chloride (CTAC) or bromide (CTAB) 1.3, 0.92

Tetradecyltrimethylammonium chloride (TTAC) bromide (TTAB) 4.5, 3.6

Hexyltrimethylammonium bromide (HTAB)

Octatrimethylammonium bromide (OTAB) 140

Propyltrimethylammonium bromide (PTAB)

Nonionic

Polyoxyethylene-23-lauryl ether (Brij-35)

Octyl-^-D-glucopyranoside (OG) 25

Nonanoyl-N-methylglucamide (MEGA 9) 19-25

Octyl-^-D-maltopyranoside 23

n-Octanoylsucrose 24

Triton X-100

A. ANIONIC SURFACTANTS

The alkyl chain can be varied to change the hydrophobicity of the formed micelles. Surfactants with alkyl chains of less than eight carbons are not very useful, since their CMCs are far too high; however, they can be used as ion-pairing reagents to modify selectivity (43-45). Alkyl chains of greater than 14 carbons pose solubility problems (46). The best alternative to SDS is sodium decyl sulfate (47), though alternatives are not generally needed.

B. CATIONIC SURFACTANTS

Cationic surfactants have the unique ability to reverse the charge of the capillary wall and, thus, of the EOF. Separations are performed using reversed polar-

4.6 Alternative Surfactant Systems 159

ity, where the negative electrode is designated as the inlet. Charge reversal occurs at surfactant concentrations well below the CMC, but without the characteristic selectivity that accompanies MECC.

Varying the size of the alkyl chain does not change the EOF, but the micel-lar mobility is modified. The longer the alkyl chain, the narrower the elution window (48).

In many cases, separations can be performed using either SDS or a cationic surfactant. For those cases, SDS is preferred, unless there are other reasons such as shortened analysis time (49). If low-UV detection is employed, the noise levels are higher with the cationic surfactant due to its higher (than SDS) UV absorption. Once a capillary has been treated with a cationic surfactant, it should not be used for any other purpose.

C. NoNiONic SURFACTANTS

Nonionic surfactants such as Brij-35 can be used to separate charged solutes (50, 51). Presumably, the only interactions between the solute and the micelle are hydrophobic, although it has been reported that nonionic micelles can adsorb ions onto its surface (52).

Alkylglycoside surfactants are neutral carbohydrate surfactants. An in situ charge in the presence of borate buffer is developed through complexation (53-57). The ratio of surfactant to borate determines the effective charge. The efficiencies appear greater than those found with conventional surfactant systems. Typical electrolytes contain 200 mM borate and 100 mM of the surfactant. Voltages are limited to 15 kV in 50-|Lim-i.d. capillaries to avoid heating problems. Surfactants such as octyl-j8-D-glucopyranoside are commercially available. These surfactants are less hydrophobic than SDS, which contributes to decreased retention of hydrophobic species.

D. BILE SALTS

Bile salts form micelles with an aggregation number of up to 10 (58). The structure and physical properties of several bile salts are given in Figure 4.13. The molecular structure of these micellar aggregates differs substantially from the long-chain alkyl variety. The hydroxyl moieties all line up in the same plane; thus, the surfactant possesses both hydrophilic and hydrophobic surfaces.

These surfactants have limited utility for chiral recognition, but they are useful for separating cationic solutes that bind strongly to SDS and for resolving hydrophobic solutes that have a migration time equal to t c- The interior of the bile salt micelle is less hydrophobic than SDS (59). Other operating characteristics such as pH and organic modifier control are similar to those of SDS, though bile salts are more tolerant of organic modifiers (59). The CMC of


COR.

BILE SALT SODIUM TAURODEOXYCHOLATE

SODIUM DEOXYCHOLATE

SODIUM TAUROCHOLATE

SODIUM CHOLATE

R i OH

OH

OH

OH

Ra H

H

OH

OH

R3 OH

OH

OH

OH

R4 NHCHaCHgSOaNa

ONa

ONa

CMC (M) 0.009 0.006

0.013

FIGURE 4.13 Structure and properties of some bile salt surfactants.

sodium chelate does not change appreciably until the methanol content is above 30%. For SDS, changes in the CMC begin at the 10% methanol level. A separation of corticosteroids is shown in Figure 4.14.

Rational selection of the appropriate bile salt is not obvious, since it not possible to predict the selectivity in advance. Bile salts have also been mixed with other surfactants to adjust the selectivity of the separation (60, 61).

E. MISCELLANEOUS SYSTEMS

Complexities and availability notwithstanding, a plethora of reagents and variants of the MECC technique abound in the literature. Polymeric cationic surfactants such as polybrene can be used for ion-exchange-like separations of acidic compounds (12, 13). The equilibrium model is similar to that presented for secondary equilibrium. Polymerized sodium undecylene sulfate has been used for separation of polycyclic aromatic hydrocarbons (62) in 25 min. These huge aggregates can be thought of having a CMC of 1 molecule. Chiral versions have been used for chiral recognition (63), but none of these are commercially available at this writing.

The first reports of the use of microemulsions for electrokinetic separations appeared in 1991 (4, 5). Microemulsions consisting of, for example, hep-tane:SDS:butanol:pH 7 buffer (0.81:1.66:6.61:90.92) can be used for separations

4.7 Cyclodextrins 161

u 12

Time (min)

16 20

FIGURE 4.14 Separation of corticosteroids with a bile salt surfactant. BGE: 100 mM borate, pH 8.45, 100 mM sodium cholate; voltage: 12.5 kV; temperature: 25°C; detection: UV, 254 nm. Key: (1) triamcinalone; (2) hydrocortisone; (3) betamethasone; (4) hydrocortisone acetate; (5) dexam-ethasone acetate; (6) triamcinalone acetonide; (7) fluocinolone acetionide; (8) fluocinonide. Reprinted with permission from the Beckman Chromatogram, August 1990.

with a utility similar to MECC (6). Of particular note is the use of such a system to indirectly determine water:octanol partition coefficients (64).

4.7 CYCLODEXTRINS

Cyclodextrins (CDs) are macrocyclic oligosaccharides that are synthesized by the bacterial enzymatic digestion of starch. The basic structures comprise six, seven, or eight glucopyranose units attached by a-1,4 linkages and are referred to as a-, /3-, and /-cyclodextrins. In addition to the native CDs, many deriva-tized cyclodextrins are now used, particularly for chiral recognition.

The interior of the CD is quite hydrophobic and is optically active. Figure 4.15 shows a view of a-CD looking down into the molecule. The shape of the


f i n (IBb tlnaQrit

woe

at 0 6 Cftrboiw >

FIGURE 4.15 Structure of a-cyclodextrin. Reprinted with permission from J. Liq. Chromatogr., 15, 961 (1992), copyright © Marcel Dekker.

molecule is cylindrical, except the diameter is tapered; this is called a torus. This is better illustrated in Figure 4.16, which also shows the nature of the inclusion complex. CDs can effectively solubilize poorly soluble solutes by formation of an inclusion complex, provided the size and shape of the compounds conform to the interior dimensions of the torus. It is also possible that a solute can sit at the opening of the CD.

The important physicochemical characteristics of CDs are listed in Table 4.3. Single-ring aromatic solutes with few side chains are best separated using a-CD. /3-CD is best for one- to two-ring aromatic compounds, whereas y-CD is used for even larger molecules. The solubility of native /J-CD is poor. The material can be solubilized with urea, but functionalized CDs such as hydrox-ypropyl-/?-CD effectively solve the solubility problem.

Most electrokinetic applications employing CDs are in chiral recognition, which we cover in Section 4.9. Cyclodextrins can be used for secondary equilibrium in achiral separations as well. For example, the addition of 2 mM dimethyl-/?-CD in 25 mM borate adjusted to pH 2.4 with phosphoric acid is used in the stability-indicating separation of the drug ranitidine from its impurities and degradants (65).


OH

***C;ir''*'—V-»w.-.'' /Estradiol

FIGURE 4.16 Possible appearance of an inclusion complex between estradiol and a cyclodextrin. Reprinted with permission from J. Liq. Chromatogr., 15, 961 (1992), copyright © Marcel Dekker.

The use of cyclodextrins follows the general principles of secondary equilibrium, which were outlined in Section 3.5. If the solutes are charged and have identical mobilities, then a neutral or charged CD can be employed. If the equilibrium constants for the formation of the inclusion complexes differ, then a separation will occur. If the solutes are neutral, then a charged CD is required. For example, CDs were used to separate structural isomers of substituted benzoic acids (66). When using a charged CD such as sulfobutylether4-j8-CD, the anionic CD countermigrates against the EOF much as the SDS micelle does. In this regard, the CD can serve as the slowly moving "phase" in electrokinetic chromatography Since micelles are so effective for this task, CDs are not as widely used, except in the area of chiral recognition.

Table 4.3 Important Characteristics of Cyclodextrins

Parameter

Molecular weight

Diameter of cavity (A)

Volume of cavity (A )

Solubihty (g/100 mL, 25°C)

Molecules per unit cell

a

972

4.7-6

176

14.5

4

Type of CD

P

1135

8

346

1.85

2

7

1297

10

510

23.2

6

Data from Luminescence Applications in Biological, Chemical, and Hydrological Sciences, ACS Symposium Series 383, p. 169.


In addition to the aforementioned achiral separation of benzoic acid structural isomers, cyclodextrins have been used in achiral applications including estrogens (67), positional isomers of methylbenzoates (68), leukotriene positional isomers (69), and ergot alkaloids (70). The BGE for the ergot alkaloids consisted of 20 mM j8-CD, 8 mM y-CD, 2 M urea, 0.3% polyvinylalcohol in phosphate buffer, pH 2.5. When used in conjunction with mass spectroscopy, concentrations of up to 20 mM did not interfere with detection (71).

Differences in the mechanism of separation with micelles or cyclodextrins account for the greater applicability of micelles in HPCE separations. The relationship between a solute and a micelle is a surface interaction. For CDs and solutes, an inclusion complex forms. This imposes steric factors, which are not found when micelles are employed.

Because of these mechanistic differences, the combination of micelles and cyclodextrins is particularly powerful, especially for nonpolar compounds such as polycyclic aromatic hydrocarbons (72) and positional isomers of nitroaro-matic compounds (73).

The separation mechanism for CD-MECC is illustrated in Figure 4.17. Micelles and CDs coexist in aqueous solution with little interaction. Underivatized CDs are neutral and have a hydrophilic outer surface, and so there is little driving force for micellar interaction. The CD in this example is simply carried by the EOF

NEUTRAL CD

ANIONIC MICELLE

EOF

INCLUSION COMPLEX

FIGURE 4.17 Separation mechanism of CD-MECC. The solute partitions between the micelle and a cyclodextrin.


toward the negative electrode. SDS micelles electromigrate toward the positive electrode as usual. Hydrophobic solutes that are normally bound to the micelle can form inclusion complexes with the CDs. The separation mechanism is then based on differences in a solute's partition coefficient between the micelle and the CD. Increasing the CD concentration will decrease ¥ for compounds that form inclusion complexes within the micelle, as shown in Figure 4.18. Whereas Figure 4.18 illustrates a decrease in k'for some corticosteroids with j3-CD, there was little change in the selectivity. The use of /-CD, shown in Figure 4.19, provides substantial improvements in corticosteroid separations. The larger cavity of the 7-CD better accommodates the bulk of the steroid moiety.

1

30 40 50 p-CD CONCENTRATION {mM)

FIGURE 4.18 Effect of j3-CD concentration on 1/k'of corticosteroids. BGE: 50 mM SDS, pH 9.0, borate-phosphate with 4.0 M urea; capillary: 50 cm length to detector x 50 jLim i.d.; voltage: 20 kV; temperature: ambient; detection: UV, 220 nm. Key: (a) hydrocortisone; (b) hydrocortisone acetate; (c) betamethasone; (d) cortisone acetate; (e) triamcinolone acetonide; (f) fluocinolone acetonide; (g) dexamethasone acetate; (h) fluosinonide. Redrawn with permission from J. Liq. Chromatogr., 14, 973 (1991), copyright © Marcel Dekker.


15 30

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FIGURE 4.19 Effect of y-CD concentration on the migration times of eight corticosteroids. The dashed hne indicates the migration of methanol, an unretained EOF marker. Other conditions as per Figure 4.18. Redrawn with permission from J. Liq. Chromatogr., 14, 973 (1991), copyright © Marcel Dekker.

4.8 APPLICATIONS AND METHODS DEVELOPMENT

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A. URINARY PORPHYRINS


Urinary porphyrins are important precursors in the biosynthetic pathway leading to hemoglobin. Various disease and toxic states interrupt the synthetic cascade, leading to a buildup of porphyrins in urine and other body tissues. These conditions are known as porphyrias. The quantitation of some of the various porphyrins are diagnostic for many of these conditions.

The structure of mesoporphyrin, a synthetic porphyrin not found in nature, is shown in Figure 4.20. Other species differ in the degree of carboxylation at the perimeter of the porphyrin ring structure. Mesoporphyrin is doubly car-boxylated, followed by coproporphyrin (4 COOHs), pentacarboxylporphyrin (5 COOHs), hexacarboxylporphyrin (6 COOHs), heptacarboxylporphyrin (7 COOHs), and uroporphyrin (8 COOHs). At the time of this work, the author was involved in the development of a fluorescence detector for HPCE. Since these compounds fluoresce strongly and have many charge states, they were considered an ideal application for this detector.

Porphyrins are usually determined by LC via gradient elution. Since the important porphyrins contain between two and eight carboxylic acid sub-stituents, they are good candidates for HPCE as well. A CZE separation is shown in Figure 4.21 and compared with one by HPLC. Isomers of hexacarboxylporphyrin not separated by LC are clearly resolved by CZE. The elution order is reversed for the two techniques. By LC, the more polar carboxylated porphyrins

VH2CH3

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FIGURE 4.21 (Main figure) CZE of urinary porphyrins. Capillary: 72 cm (50 cm to detector) x 50 ]im i.d.; BGE: 20 mM CAPS, pH 11, with 10% methanol; sample porphyrin test mix, 5 nmol/mL in methanol:20 mM CAPS (50:50); injection: electrokinetic, 12 s at 10 kV; voltage: 30 kV; detection: fluorescence; excitation: 400 nm; emission wavelengths > 595 nm. (Inset) Gradient elution reversed-phase LC of urinary porphyrins. Key: (1) mesoporphyrin (dicarboxyl); (2) copropor-phyrin (tetracarboxyl); (3) pentacarboxyl porphyrin; (4) hexacarboxylporphyrin positional isomers; (5) heptacarboxyl porphyrin; (6) uroporphyrin (octacarboxyl). Reprinted with permission from J. Chromatogr., 516, 271 (1991), copyright © 1991 Elsevier Science Publishers.

elute first. By CZE, the most highly charged porphyrins migrate toward the positive electrode but are swept toward the negative electrode by the EOF. In this case, the more polar and charged species elute last.

Although the CZE separation appears adequate, repeated runs show a merging and broadening of peaks, characteristic of solutes binding to the capillary walls. Since the porphyrins are all anionic, mesoporphyrin, which is poorly soluble, binds through hydrophobic interaction of the uncharged quadrant of the

molecule with the capillary wall. This is indicated in Figure 4.21 by the short broad band surrounding peak 1.

The porphyrins are anionic and hydrophobic, and thus MECC seemed appropriate, since SDS is anionic and hydrophobic as well, thereby increasing the likelihood that the active sites on the capillary wall would be saturated. The mechanism of separation would be expected to resemble the separation without surfactant, since the anionic porphyrins would be expected to be repelled from the anionic micelle.

In 100 mM SDS, pH 11 (Figure 4.22A), the elution order is the same as CZE except for mesoporphyrin. Mesoporphyrin has its carboxylate groups located on one quadrant of the molecule. The other side of the molecule is free to interact hydrophobicly with the micelle. At 150 mM SDS (Figure 4.22B), the mesoporphyrin exhibits a further shift in migration time that is consistent with this argument.

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TIME(min.) FIGURE 4.22 MECC of urinary porphyrins: Capillary: 77 cm (55 cm to detector) x 50 |lm i.d.; sample: porphyrin test mix, 20 nmol/mL dissolved in 20 mM CAPS, pH 11, 100 mM SDS; (A) run BGE: 20 mM Caps, pH 11, 100 mM SDS; voltage: 20 kV; temperature 45°C; (B) run BGE: 20 mM Caps, pH 11, 150 mM SDS; voltage 20 kV; temperature 45°C. Key: see Figure 4.21. Reprinted with permission from J. Chromatogr., 516, 271 (1991), copyright © 1991 Elsevier Science Publishers.

For coproporphyrin, peak 2 exhibits fronting. This is due to a solubihty problem, wherein it is not very soluble in the bulk solution nor in the micelle. An organic modifier was employed to solve the solubility problem (Figure 4.23). With 15% methanol, a sharper peak is obtained, but the time of separa-

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tion is prolonged due to a reduction in the EOF Acetonitrile would have been a better choice of modifier, since it does not reduce the EOF To speed the separation, both increased temperature and increased voltage were applied. At 30 kV, 45"^C, the time of separation is only 13 min, and the coproporphyrin peak is now very sharp.

Next, a urine sample taken from a patient suffering from porphyria cutanea tarda, a genetic disorder, was run (Figure 4.24). Characteristic of this disease is

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FIGURE 4.24 (Main figure) Electropherogram of a urine sample from a patient with porphyria cutanea tarda. Sample preparation: centrifugation of urine for 1 min; injection: 2 s vacuum (10 nL). (Inset) Electropherogram of partially decomposed porphyrins in a urine sample. Other conditions given in Figure 4.22A. Key: see Figure 4.21. Reprinted with permission from J. Chromatogr., 516, 271 (1991), copyright © 1991 Elsevier Science Publishers.

an elevation of peaks 5 and 6. Splitting of these peaks was noted. Perhaps the porphyrins were further separated into various classes. Standards of type I and type III isomers were studied, and it was found that separation was not occurring. However, a photodegraded standard showed the same splitting pattern, and that is why all porphyrin labs are kept dark. The splitting in the sample was due to photodegradation during a 24-h urine collection.

To complete this story, the type I and type III isomers of copro- and uroporphyrin were separated by affinity capillary electrophoresis (ACE) using an electrolyte containing 20 mM phosphate, pH 1.6, and 0.03 mM bovine serum albumin (129). Bile salt micelles also served to fractionate the isomers. A BGE containing 60 mM deoxycholic acid, 15% acetonitrile, and 10 mM borate was effective for this purpose (130). As will be described later, this porphyrin application has a few more surprises in store.

B. DRUG SEIZURE SAMPLES

HPLC is a superb technique for small-molecule separations. Figure 4.25 shows the HPLC separation of a series of drug standards that are used for screening of seizure samples by the U.S. Drug Enforcement Administration (DEA) (36). The separation employs a complex gradient and uses hexylamine to cover free silanols. The separation time is nearly 1 h including the gradient reequilibra-tion step. Fine-tuning this separation took many weeks.

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In 1990, the DEA decided to attempt this separation by MECC. The rationale for using MECC was to apply the method to acidic, basic, and neutral drug substances simultaneously. During the course of an afternoon's work, the separation shown in Figure 4.26 was developed (36). The separation was not optimized, but the run time was much shorter than for HPLC. Unlike the HPLC separation, where resolution was barely adequate, the MECC separation time could be shortened by reducing the capillary length and using a lower concentration of SDS. That separation is shown in Figure 4.27. Including the reequili-bration time, the MECC separation is now 10 times as fast as HPLC. The same BGE can be used for screening cocaine samples. For screening of drug seizure samples, a single CE unit could replace 10 liquid chromatographs.

C. A BASIS FOR METHODS DEVELOPMENT

AND O P T I M I Z A T I O N

The remarkable ability of electrokinetic chromatography to provide a few hundred thousand theoretical plates permits complex separations to be performed.

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frequently without the need to employ chemometrics-based optimization schemes. Following a scheme such as that illustrated in Figure 4.28 will usually lead to an adequate separation, often during the course of less than a day of experimentation. More often than not, the 100 mM SDS, 20 mM borate buffer will yield a good starting point.

Switching to surfactants other than SDS is normally beneficial for the separation of nonpolar substances or when SDS alone gives too much or too little retention. Even in the former case, the use of cyclodextrins permits the separation of nonpolar species such as aromatic hydrocarbons. Usually, alternative surfactants should be considered only after other experiments covering pH, modifiers, and so forth have been performed using SDS—unless, of course, a suitable reference has been located. Most problems will be solvable using SDS or SDS with various additives. Even if a publication reported on an alternative surfactant system, the separation may be possible with SDS. Often, several different surfactant systems are suitable for a given problem.

On the other hand, bile salts are useful for separating rigid planar molecules such as steroids (131). The sterol architecture resembles the steroid structure; thus, the old adage from freshman chemistry, "like dissolves like," applies here. In a similar fashion, the planar macrolide antibiotics are well separated using


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bile salt surfactants (105). In any event, a few scouting runs should point you in the correct direction.

Once the appropriate system has been identified through scouting runs, optimization is usually straightforward. Adjustments in pH and additive concentrations should be carefully studied. It is best not to rely on a long capillary to perform the separation, although this may become necessary when many components are being resolved. In complex samples, solving the separation of a pair of overlapping components often causes coelution of other solutes.

For these situations, statistical tools can be a valuable tool for speeding methods development. This can be particularly true when ternary blends of solvents or cyclodextins are needed to optimize the separation. For example, overlapping resolution maps (ORM) have been used for years to optimize HPLC separations. In 1991, this technique was used to optimize the separation of plant growth regulators using mixtures of a-, /?-, and /-CD (132), and in 1997, it was applied to optimize the separation of quinolone antibacterials using cholate and heptane sulfonate (133). A pure chemometric approach using Plackett-Burman statistics has also been shown useful in the separation of testosterone esters (134). While these tools are not often used, they should be considered when trial and error proves frustrating.

4.9 Chiral Recognition 1 7 9

4.9 CHIRAL RECOGNITION

A. BASIC CONCEPTS

Chiral recognition of racemic mixtures continues to be an active area of research in gas chromatography, hquid chromatography, and of late, capillary electrophoresis. Whatever the separation technique employed, chiral recognition is obtained in one of three ways:

1. Formation of diastereomers^ by additives to the mobile phase or carrier electrolyte

2. Formation of diastereomers through interaction with a stationary phase or the functionalized capillary wall

3. Precolumn (capillary) derivatization with an optically pure derivatizing reagent

In the first two cases, diastereomer formation is transient, occurring via electrostatic and/or hydrophobic mechanisms. Since derivatization is not employed, the enantiomers are directly separated. In the third case, covalently bound derivatives are separated by MECC. Derivatization is advantageous when the solute lacks a good chromophore. In this case, the problem of chiral recognition and that of detection sensitivity both are solved in a single step.

Chiral recognition in HPCE employs secondary equilibrium for the separation of enantiomers. Reagents such as cyclodextrins, bile salts, mixed micellar systems with chiral surfactants, crown ethers, macrocyclic antibiotics, proteins, heparins, dextrins, oligosaccharides and other carbohydrates, and trimolecular peptide-Cu(II)-amino acid complexes and MECC resolution of preformed diastereomers have all been reported. Polymerized surfactants and cyclodextrins have also been used, but these are not currently available. Other unusual reagents for chiral recognition appear in the literature as well. A summary of applications and buffer recipes is given in Table 4.5. HPCE is distinctly superior to LC for chiral recognition in cost, speed, and resolution. Chiral columns for LC are relatively expensive.

B. METAL-ION COMPLEXES

The first examples of chiral recognition in HPCE employed the addition of Cu(II) and L-histidine to the buffer solution to resolve dansyl amino acids via a trimolecular complex (135). A Cu(II)-aspartame complex was later shown to be superior (47). Addition of a surfactant can improve the hydrophobic aspects of the separation, permitting the simultaneous resolution 14 out of 18 dansyl

^Diastereomers (also called diastereoisomers) are stereoisomers that contain at least two asymmetric centers that are not mirror images of each other.

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n

3


amino acids (47). The use of 2.5 mM CuS04»5H20, 5.0 mM aspartame, and 10 mM ammonium acetate, pH 7.2, gave the best separation. Addition of 20 mM STS provided hydrophobic selectivity via the MECC mechanism. The capillary had to be rigorously conditioned with 100 mM phosphoric acid for several hours to remove any metal hydroxides that may have been precipitated at the capillary wall. After rinsing for 10 min with KOH, the capillary was conditioned in acetate buffer for 10 h. Amino acid separations are the only known applications of this technique.

C. CHIRAL SURFACTANTS AND MIXED MICELLES

Separations have been reported employing a mixture of SDS and optically active surfactants such as N,N-dodecyl-L-alanine [in the presence of Cu(II)] or sodium N-dodecanoyl-L-valinate (SDVal) (17, 173). Separations reported to date have been limited to derivatized amino acids.

Mixtures of SDS with the nonionic optically active surfactant digitonin (140) have been reported to separate PTH-amino acids. A low buffer pH was selected so that electrophoresis greatly dominated electroendosmosis. The separation times were lengthy, and the early eluting solutes were not adequately resolved.

Surfactants specifically designed for chiral separation are commercially avail-able^ (148, 174). The surfactants are either of the enantiomers of N-dodecoxy-carbonylvaline. By selection of the alternate enantiomer, the order of elution of the chiral solutes can be changed. This is advantageous, since it is often desirable to have the trace enantiomer elute first.

Separations are scouted and optimized in a similar fashion as in conventional MECC. For hydrophobic neutral solutes and bases, 25 mM surfactant concentration in 50 mM borate is usually sufficient. Acetonitrile can be added if the solutes elute near t^^. For acidic solutes that are repelled from the anionic micelle or for hydrophilic neutrals and bases, the surfactant concentration can be raised to 100 mM. The surfactants are not soluble under acidic conditions.

D. CYCLODEXTRINS

Cyclodextrins (CDs) are the reagents of first choice for chiral recognition. Results such as that illustrated in Figure 4.29 are frequent occurrences. The vast majority of the reported HPCE chiral separations employ these compounds. The first HPCE reports appeared in 1988. In one article, the cyclodextrin was incorporated in a rigid gel (19); in another, HPCE with CDs was used for isota-chophoretic separations (175). The following year, the use of CDs as electrolyte additives in chiral CZE appeared (18), although CDs were reported for achiral

^Enantioselect, Scientific Resources Inc., Eatontown, NJ.

4.9 Chiral Recognition 183

10 o CM

^ u 1 ^ 2 4 mm.

FIGURE 4.29 Separation of (+) and (-) terbutaline with 5 mM heptakis(2,6-di-0-methyl)-)8-CD. Capillary: polyacrylamide-coated 20 cm x 25 liim i.d.; BGE: 100 mM phosphate, pH 2.5; constant current: 19 (xA; voltage: 9 kV; injection: electrokinetic, 8 ky 10 s of a 10~5 M solution. Reprinted with permission from J. Chromatogr., 545,437 (1991), copyright © 1991 Elsevier Science Publishers.

separations as early as 1985 (9). Since that time, hundreds of papers have appeared in the hterature.

The first generation of cyclodextrins were the native a-, j8-, and y-CD. Because of the relatively low solubility of j3-CD and the inability of native CDs to resolve all enantiomers, it was not long before CD derivatives were used in HPCE (176-178). The next generation were the negatively charged sul-fobutylether-/3-CDs (179).^ This CD is charged at all pH values and, like SDS, countermigrates against the EOF. The fourth generation are the highly substituted single-isomer CDs (180,181). These materials are well characterized and prevent the possibility of the separation being affected by the isomer content of the CD. These and similar highly sulfated CDs are commercially available.^ Because of the high currents that are generated, a 25-|am-i.d. capillary is generally required when highly charged CDs are used, but their resolving power is usually superior to other reagents. They are also quite expensive.

"^Available from Cydex, Overlook Park, KS, and Bioscience Innovations, Lawrence, KS. 5Regis Chemical, Morton Grove, IL, and Beckman Instruments, Fullerton, CA.


Chiral recognition kits and protocols are available from a number of instrument manufacturers, including Hewlett-Packard and Beckman Instruments. Cyclolab, a company located in Budapest, Hungary, specializes in CDs and offers many different derivatives of all three native materials.

The mechanism of chiral resolution is based on differences in the complex stability between each enantiomer and the CD. The basic principles of secondary equilibrium apply here. The Wren and Rowe model, which is generally applicable to secondary equilibrium, was originally developed for chiral recognition with CDs (182-184). If the solutes spend either too little or too much time attached to the CD, no separation occurs. Then, the CD concentration or type must be changed. For solutes that are too strongly bound, the addition of organic solvents can be helpful. Both hydrophobic inclusion within the CD and hydrogen bonding between the analyte and CD hydroxyl groups are in part responsible for resolution. The molecular fit of the solute within the CD cavity is critical if chiral recognition is to be obtained. It is generally not possible to predict if a separation will occur.

More sophisticated models have recently appeared in the literature that seek to explain the relationships of pH and CD concentration on chiral separations. The models also account for the reversal of elution order at different pHs (185-187). While not covered here, elegant 3-D graphs provide an understanding for the development of optimal conditions.

The relatively long history of LC and GC provides a framework for determining if applications will be successful. Armstrong's empirical procedure employs molecular structure to predict enantioselectivity in LC using a func-tionalized/J-CD (188). In this model, sp^-hybridized carbons connected to the stereogenic center were found to provide enhanced resolution. Conversely, sp^-hybridized carbons showed diminished stereoselectivity. Amido groups improved selectivity, especially when associated with ;r-acid (3,5-dinitroben-zoyl) or ;r-basic (naphthyl) groups. These studies employed 121 compounds to develop the model.

Most of the reported literature on CDs employs CZE on uncoated capillaries. It is possible to immobilize CDs in a gel (19) or use a coated capillary with immobilized CD (189-191). The gel format lowers the EOF and improves resolution, but the inconvenience of gel-filled capillaries is substantial. In the latter case, open tubular capillary electrochromatography is being performed. Because of mass transport problems, the efficiencies are generally less than those found in CZE. If it is necessary to lower the EOF, a polymer network (176) or coated capillary is a simpler solution. Figure 4.30 shows the separation of chloramphenicol at pH 3.5 with and without the polymer network. It is possible that by lowering the pH to 2.5, the reduction of EOF would eliminate the need for the network.

Charged solutes can be separated using neutral CDs. Adding the appropriate CD to 50 mM phosphate buffer, pH 2.5, for bases and 50 mM borate buffer, pH


FIGURE 4.30 Influence of methylhydroxyproplycellulose on the chiral recognition of chloramphenicol enantiomers (A) without methylhydroxyethylcellulose (MHEC) and with 0.1% MHEC. BGE: 20 mM Tris adjusted to pH 3.5 with citric acid with 10 mM heptakis(2,6-di-0-methyl)-j8-CD; capillary: 65 cm (45 cm to detector); voltage: 18 kV; current: 6 |LIA; detection: UV, 254 nm. Reprinted with permission from J. Chromatogr., 559, 215 (1991), copyright © 1991 Elsevier Science Publishers.

9.3, for acids are good starting points. Use CD concentrations of 5 and 20 mM. If no separation occurs, first adjust the pH to equal the pK^ of the solute. If the separation still does not occur, try another CD. For separating neutral compounds, charged CDs must be employed.

Once scouting runs have revealed a separation, optimization is relatively straightforward. It is important during these scouting runs that the direction of migration be determined. This is particularly important when using charged CDs at low pH, where the EOF is minimal. The direction of migration is then


determined by the strength of the solute-CD complex (192). In this example, where mixed CDs are employed, a solute such as cocaine that has a high affinity for the anionic CD migrates toward the anode. Solutes such as amphetamines have a low affinity for the anionic CD and migrate toward the cathode. Some scouting runs on the short end of the capillary ensure that the proper polarity of the power supply is selected.

When charged cyclodextrins are used for separating charged solutes, ion pairing or ion repulsion can occur (178). In this case, reversal of the enantiomer elution order can occur as well, and depending on the strength of the solute-CD interaction, migration can occur in either direction, particularly at low pH or when coated capillaries are used as described in the previous paragraph. When necessary, ion-pairing reagents can be added to the BGE to improve separations. Such reagents include quinine (193), tartaric acid (194), and various sulfonic acids (195).

Once the appropriate CD is selected, the important variables to optimize are the CD concentration, the buffer concentration, and the temperature. If the resolution is optimized, then the benefits of a shortened capillary length can be enjoyed.

Most solutes have relatively weak affinity for the CD. This is illustrated using the drug quinagolide as an example in Figure 4.31 (top) (20). As the buffer concentration is increased, the resolution of the enantiomers increases in a nearly linear manner until the point where Joule heating becomes important. A concentration of 50 mM was optimal under the conditions studied. Since Joule heating can be limiting, decreasing the capillary diameter and/or the field strength might further improve the resolution by permitting the use of higher concentration buffers. The buffer type also played a role in chiral resolution. Glycine HCl, pH 2.5, 50 mM gave poorer resolution than phosphate, pH 2.5, and no resolution was found in 50 mM citrate, pH 2.5.

A linear dependence of the migration time on the concentration of/J-CD was found in the concentration range 10-30 mM. No separation occurred below 10 mM CD concentration. This is consistent with the solute's weak interaction with the CD. If the interaction were strong, the resolution would begin to decline as the equilibrium was pushed too far to the right.

The impact of temperature is substantial in chiral HPCE. As Figure 4.31 (bottom) indicates, lowering the temperature improves the resolution. This will usually be the case when there are weak interactions between the solute and the CD.

The order of elution of the enantiomers can be important. Drug purity determinations entail the undesirable enantiomer to be present at quantities of less than 0.1% relative to the major component. For these separations, it may be advantageous for the impurity to elute prior to the main component. For example, a peptide drug with two chiral centers, separated at pH 2.5 with sul-fobutylethery-jS-CD, has the trace impurity eluting after the major component.

4.9 Chiral Recognition

1.2 y

1.0 4-

0.8 4-

R 0.6-1-

0.4-|-

0.2 4-

187

0.0 4- 4- 4-0.00 0.02 0.04 0.06 0.08

Phosphate cone. |nK>l t'^ ]

0.10

1.2-y

1.0 4-

0.84-

R 0.6 4-

0.44-

0.24-

0.0 15 20 25 30 35 40 45

Temperature fC] FIGURE 4.31 Influence of BGE concentration (top) and temperature (bottom) on the resolution of the enantiomers of quinagolide. Capillary: 50 cm X 75 |lm i.d.; voltage: 15 kV; BGE: 50 mM phosphate, pH 2.5, p-CD (top, variable; bottom, 50 mM); detection: UV, 214 nm. Reprinted with permission from Chromatographia, 33, 32 (1992), copyright © 1992 Vieweg.


By running at pH 9.3, the order of migration switches, and the trace component elutes first. Such a separation is shown in Figure 4.32.

The separation shown in Figure 4.32 deserves further comment. A long 108-cm capillary was used here with an electrolyte containing 50 mM borate and 20 mM sulfobutylethery-^-CD. The current was 70 |iA at 30 kV, 30°C, using a 50-|im-i.d. capillary. Reducing the temperature to 15^C reduced the current, permitting a shorter capillary to be employed at 30 kV The run time decreased to 40 min with baseline resolution.

E. CROWN ETHERS

Crown ethers, represented in Figure 4.33 by 18-crown-6 tetracarboxylic acid, are another class of complexing reagents that have been employed in LC for

TIME (min)

FIGURE 4.32 Separation of trace enantiomers in the presence of the major component: Capillary: 108 cm X 50 |xm i.d., "bubble factor 3"; BGE: 20 mM sulfobutylethery-jS-CD, 50 mM borate, pH 9.3; injection: 100 mbs; detection: UV, 200 nm; voltage: 30 kV; temperature: 30 °C; solute concentration: 1 mg/mL.

COOH^^,^ O \ ,^>COOH

COOH^< \ O' COOH

FIGURE 4.33 Structure of 18-crown-6 tetracarboxylic acid.

chiral recognition. This reagent is useful for chiral separation of primary amines (20, 196, 197). The material is no longer commercially available, and it was very expensive. It was shown in 1997 and 1998 that by combining CDs in conjunction with inexpensive nonfunctionalized 18-crown-6, comparable separations could be obtained (164,198,199). Typical BGE recipes include 50 mM phosphate, pH 2, 5 mM 18-crown-6 and either 5 mM dimethyl-j8-CD or 20 mM 7-CD, depending on the application.

F. MACROCYCLIC ANTIBIOTICS

Macrocyclic antibiotics are a broad class of naturally occurring compounds that have numerous chiral centers and a variety of functional groups that are known to produce chiral separations (200). The structure of one such compound, vancomycin, is shown in Figure 4.34. Macrocyclics such as vancomycin (200), teicoplanin (143), and rifamycin B (11) have all produced chiral separations. Ristocetin A may be the most widely applicable of these glycopeptide compounds (201). While there is certainly overlap within the scope of compounds separated by CDs, the chiral resolving power of these selectors may be superior.

These selectors are not without difficulties. The main disadvantage is strong absorption of light in the mid- to low-UV region of the spectrum. This requires that the solute not be detected at wavelengths where the antibiotic absorbs. For solutes lacking a chromophore, they can be used simultaneously for both indirect detection and chiral recognition when a solute does not absorb light (11).


Vancomycin R = H

Me, ^NHj

A82846B R

FIGURE 4.34 Structure of a macrocyclic antibiotic. Courtesy of John Reilly, Eli Lilly and Company Limited, United Kingdom.

Typical electrolytes contain 40-100 mM phosphate buffer, pH 6, and 0.5-5 mM of the antibiotic. Organic solvents such as acetonitrile may be necessary additives when solute binding to the antibiotic is very strong (143). The solvent 2-methoxyethanol was shown more effective than acetonitrile for resolving nonsteroidal anti-inflammatory drugs (202).

If the solute's UV absorption spectrum overlaps the that of the antibiotic, it may be possible to overcome this problem via a special technique (203). A coated capillary is used to suppress the EOF As Figure 4.35 indicates, the capillary is filled with BGE containing the antibiotic, and a sample is loaded onto the capillary. Operating in the reversed-polarity mode (sample-side negative), the cationic antibiotic migrates toward the negative electrode away from the detector, while the anionic solute migrates toward the positive electrode. Since the outlet electrolyte reservoir does not contain the antibiotic, the detector window is clear when the separated enantiomers reach it. Naturally, this system works only for anionic solutes. For cationic solutes, an anionic antibiotic would be required. This might involve adjusting the pH, since many of the macro-cyclics are zwitterionic. A separation of dansyl glutamic acid using this approach is shown in Figure 4.36. Note the rapidly stabilizing baseline as the antibiotic clears the detector.


^^,,.^...^^^^^^^^^,^^^^^^^2 ^^-'^^^^^^^^^^^^^'^^^^^s^^^^Mm^^

FIGURE 4.35 Schematic illustrating the separation of an ionic drug using a macrocyclic antibiotic. The UV-absorbing macrolide is not present in the electrolyte reservoirs. A coated capillary and reversed polarity are used to ensure the capillary window is clear of the antibiotic prior to detection of the solutes.

G. OLIGOSACCHARIDES

Oligosaccharides comprise a vast array of compounds that include linear ohgosaccharides, maltodextrin mixtures, maltooligosaccharides, linear non-a-(l-4)-linked glucose polymers, low-molecular-mass galactose-glucose-fructose copolymers, and even cyclodextrins (145). These compounds and others such as amylodextrins can form complexes with many small molecules (204).

A maltooligosaccharide such as 10% Dextrin 10 solution in 100 mM phos-phate/pyrophosphate, pH 7 was more effective than 10% Dextran or 20 mM /3-CD in separating some nonsteroidal anti-inflammatory drugs (148). It is known that derivatized CDs are often required for chiral recognition of these classes of acidic drugs (205). It has been found that a degree of polymerization (DP) greater than 7 was required for chiral recognition (172).

Separations of a variety of drug substances have been demonstrated with 3% chondroitin sulfate C, a mucopolysaccharide, in 20 mM borate-phosphate, pH 2.8. That reagent was favored over heparin sulfate and dextran as a chiral selector (206).


875

843

400

Time (Seconds)

FIGURE 4.36 Chiral separation of dansyl glutamic acid. Conditions: capillary: 27 cm (20 cm to detector) x 50 m eCAP neutral (Beckman); BGE: 2 mM vancomycin, 100 mM phosphate, pH 6; the voltage was applied for 5 min prior to injection to purge the detection window of antibiotic; injection: 2 s, low pressure (0.5 psi); detection: UV, 254 nm; voltage: -10 kV; temperature: 25°C; sample concentration: 0.1 mg/mL. Courtesy of John Reilly Eli Lilly and Company Limited, United Kingdom.

The advantages and disadvantages of these materials compared with cyclodextrins are unclear at this time. Only a dozen papers have been published, and the range of compounds studied overlaps with those studied with CDs.

H. BILE SALTS

Bile salts have already been shown useful for the determination of hydrophobic solutes. This usefulness can be extended to the separation of optical isomers; bile salts are naturally optically active. Unlike the work with CDs, bile salts are best utilized under conditions of a strong EOF; this is a consequence of the separation chemistry following the MECC mechanism. Operation at low pH values generally results in lengthy separation times.

Bile salts tend to provide enantioselectivity when the structure of the solute is more rigid than that of the surfactant (146). The selection of the appropriate bile salt has a substantial impact on the chiral recognition. There is no way of predicting which salt will yield the best results at this time. High buffer pH values where solutes become anionic reduce chiral recognition, presumably due to repulsion from the anionic micelle (146). Neutral compounds are not affected by pH changes. The use of organic solvents to reduce K'and improve a has been reported (207). A textbook separation is shown in Figure 4.37.


U n Hi if

U — I I L —r-19

"""""I -'l

12 14 —r-16 18 20

FIGURE 4.37 Chiral separation of trimetoquinol HCl, tetrahydropapaveroline, five diltiazen-related compounds, 2,2'-dihydroxy-l,r-dinaphthyl, and 2,2,2-trifluoro-l-(9-anthryl)ethanol. Capillary: 65 cm (50 cm to detector) x 50 )im i.d.; BGE: 50 mM STDC in 20 mM phosphate-borate, pH 4.0; voltage: 20 kV; detection: Uy 210 nm. Reprinted with permission from J. Chromatogr., 515, 223 (1990), copyright © 1990 Elsevier Science Publishers.

Admixtures of bile salts with cyclodextrins (208) and polyoxyethylene ethers (60) have also been reported. Since very few papers have been published in this field and general utility has not been demonstrated, bile salts are usually tried only upon failure of other methods.

I. PRECAPILLARY DERIVATIZATION

Separations have been reported by MECC for amino acids derivatized with Mar-fey's reagent (l-fluoro-2,4-dinitrophenyl-5-L-alanine amide) (209), GITC (2,3,4,6-tetra-0-acetyl-j8-D-glucopyranosylisothiocyanate) (21), and l-(9-fluorenyl)-ethyl chloroformate (FLEC) (210). The GITC derivatives of amphetamine andmetham-phetamine separated with 80 mM SDS, 8 mM phosphate-borate, pH 9; 20% methanol allows simultaneous separation of the precursors as well (211).

The FLEC reagent is particularly notable. This reagent reacts rapidly with primary and secondary amines to form stable fluorescent derivatives. Detection


can be by absorption at 260 nm or laser-induced fluorescence using a krypton fluoride laser with fluorescence collected above 300 nm. The reagent meets all of the standard criteria for an ideal reagent (to be listed presently). Its only shortcoming is the need to perform a solvent extraction to remove the excess reagent. This can result in recovery losses for very hydrophobic solutes.

A variety of reagents have been employed to create synthetic diastereomers that can be separated by LC (212). It is likely that many of these will be applicable to separation by MECC. When a reagent is available in both optically active forms, it is possible to control the order of elution of the enantiomers. Ideally, the enantiomer present in the lower concentration should elute first. It is less likely to be lost on the tail of the more concentrated solute.

The ideal reagent will have the following characteristics:

1. Rapid reaction 2. Stable products 3. No racemization 4. Excess reagent invisible to detector or easily removable 5. Reagent contains or produces a strong chromophore or fluorophore 6. Commercially available in either form of purified enantiomer 7. Inexpensive

The advantages of precapillary derivatization are

1. It provides a highly predictable mechanism for chiral recognition, provided the chiral center is in the proximity of the reaction site.

2. It simultaneously produces a good chromophore and chiral selectivity.

The disadvantages of precapillary derivatization are

1. It complicates assay validation. 2. Incomplete reactions are possible. 3. Excess reagent can complicate separations. 4. There are extra sample-handling steps.

Many analytical chemists are averse to using derivatization procedures to improve separation and detection. This is unfortunate, since the enhanced results often justify the extra work in sample preparation and method validation. In chiral separations, it is possible to solve both separation and detection problems using a single procedure.

4.10 AFFINITY CAPILLARY ELECTROPHORESIS

The use of HPCE to study molecular interactions is known as affinity capillary electrophoresis (ACE). While all forms of secondary equilibrium, including the use of micelles and cyclodextrins, can be considered affinity interactions, the term ACE is generally reserved for the determination of noncovalent interactions of biomol-

4.10 Affinity Capillary Electrophoresis 195

ecules with various reagents. For example, the separation of immune complexes would be termed immunoaffinity CE (213). The study of protein-drug interactions is particularly important, since it is important for the pharmacologist to know the concentration of free drug in blood serum. The screening of combinatorial libraries for activity (binding) is another potential application (214).

ACE relates changes in the mobility of a protein (P) with a ligand (L). Either the protein or ligand can be present in the buffer, while the other is the injected solute. Analysis of the change in mobility of the protein as a function of the ligand concentration provides the binding constant K . In the absence of EOF, the change in migration time of L is proportional to the change in mobility of L (215). Then, the binding constant can be estimated by

[L] K .Ar^ -KbAtL , (4.6)

Table 4.6 lists applications that employ ACE. Affinity interactions can be precapillary (216), on-capillary (gel) (217), or postcapillary (218). The affinity

Table 4.6 Application of Affinity Capillary Electrophoresis

Biomolecule

Actinividin

ai-Acid glycoprotein

Amyeloid P component

Antithrombin

Bovine serum albumin

Carbonic anhydrase

Concanavalin A

Glycosoaminoglycans (GAGs)

Human myeloma IgE

Human serum albumin

Humic substances

Immunoglobulin G

Monoclonal anti-phosphotyrosine

Oligodeoxynucleotides

Peptides

Ligand

Biotin

Remoxipride

Heparin

Heparin

Leucovorin

Coproporphyrin

Benzenesulfonamides

Arylsulfonamides

Monosaccharides

Heparin binding peptides

DNA aptamer

Benzoin

Triazine

Protein A

Phospho to tyrosine

Poly(9-vinyladenine)

Vancomycin

Note

Chiral

Chiral

Type I and III isomers

EOF compensated

LIF

Immobilized GAG

FITC tagged

Chiral

FITC tagged

Immuno CE

Gel

Multipeptide separation

Reference

221

222

223

224

225

129

215

226

227

228

229

230

231

232

233

234

235,236


interaction can be used to effect a separation without the determination of binding constants (129). In this case, the rules of secondary equihbrium apply as always.

Depending on the reaction kinetics between the protein and ligand, some unusual peak shapes can be obtained. Some of these are illustrated in Figure 4.38. There are also many methods for performing ACE, and the merits and limitations of each have been published (219, 220).

REACTION KINETICS

Ref. It I

No interaction

Fast

Slow ^

Slow i Slow

_ ^ ^ FIGURE 4.38 Illustration of some possible patterns in affinity CE. The reference component is a noninteracting solute. Reprinted with permission from J. Chromatogr., A, 680, 405 (1994), copyright © 1994 Elsevier Science Publishers.

References 197

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158. Otsuka, K., Smith, C. J., Grainger, J., Barr,J. R., Patterson Jr., D. G., Tanaka, N., Terabe, S. Stereoselective Separation and Detection of Phenoxy Acid Herbicide Enantiomers by Cyclodextrin-Modified Capillary Zone Electrophoresis-Electrospray Ionization Mass Spectrometry/ Chromatogr, A, 1998; 817:75.

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160. Shibukawa, A., Lloyd, D. K., Wainer, I. W. Simultaneous Chiral Separation of Leucovorin and Its Major Metabolite 5-Methyltetrahydrofolate by Capillary Electrophoresis Using Cyclodextrins as Chiral Selectors: Estimation of the Formation Constant and Mobihty of the Solute-Cyclodextrin Complexes. Chromatographia, 1993; 35:419.

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162. Fanah, S., Camera, E. Use of Cyclodextrins in the Capillary Electrophoretic Separation of Ery-thro- and Threo-Mefloquine Enantiomers. J. Chromatogr, A, 1996; 745:17.

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165. Schmitt, T., Engelhardt, H. Derivatized Cyclodextrins for the Separation of Enantiomers in Capillary Electrophoresis. J. High Res. Chromatogr, 1993; 16:525.

166. Fillet, M., Bechet, I., Chiap, P, Hubert, R, Crommen, J. Enantiomeric Purity Determination of Propranolol by Cyclodextrin-Modified Capillary Electrophoresis. J. Chromatogr, A, 1995; 717:203.

167. Bjornsdottir, I., Hansen, S. H., Terabe, S. Chiral Separation in Non-aqueous Media by Capillary Electrophoresis Using the Ion-Pair Principle. J. Chromatogr, A, 1996; 745:37.

168. Nouadje, G., Nertz, M., Courderc, E Study of the Racemization of L-Serine by Cyclodextrim-Modified Micellar Electrokinetic Chromatography and Laser-Induced Fluorescence Detection. J. Chromatogr, A, 1995; 716:331.

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185. Rawjee, Y. Y., Williams, R. L., Vigh, G. Capillary Electrophoretic Chiral Separations Using p-Cyclodextrin as Resolving Agent. II. Bases: Chiral Selectivity as a Function of pH and the Concentration of B-Cyclodextrin. J. Chromatogr, 1993; 652:233.

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187. Williams, B. A., Vigh, G. Dry Look of the CHARM (Charged Resolving Agent Migration) Model of Enantiomer Separations by Capillary Electrophoresis. J. Chromatogr, A, 1997; 777:295.

188. Berthod, A., Chang, S.-C, Armstrong, D. W Empirical Procedure That Uses Molecular Structure to Predict Enantioselectivity of Chiral Stationary Phases. Anal. Chem., 1994; 64:395.


189. Armstrong, D. W, Tang, Y., Ward, T., Nichols, M. Derivatized Cyclodextrins Immobilized on Fused-Silica Capillaries for Enantiomeric Separations via Capillary Electrophoresis, Gas Chromatography, or Supercritical Fluid Chromatography. Anal. Chem., 1993; 65:1114.

190. Mayer, S., Schurig, V Enantiomer Separation by Electrochromatography on Capillaries Coated with Chirasil-Dex. HRC & CC, 1992; 15:129.

191. Mayer, S., Schurig, V Enantiomer Separation by Electrochromatography in Open Tubular Columns Coated with CHIRASIL-DEX. J. Liq. Chromatogr., 1993; 16:915.

192. Lurie, 1. S., Klein, R. X., Lebelle, M., Brenneisen, R., Weinberger, R. Chiral Resolution of Drugs of Forensic Interest by Capillary Electrophoresis Using Cyclodextrins. Anal Chem., 1994; 66:4019.

193. Thomas, J., Bunke, A., Karbaum, A. Use of Chiral and Achiral Ion-Pairing Reagents in Combination with Cyclodextrins in Capillary Electrophoresis. J. Chromatogr., A, 1998; 798:281.

194. Zhou, L., Trubig, J., Dovletoglou, A., Locke, D. C. Enantiomeric Separation of the Novel Growth Hormone Secretagogue MK-0677 by Capillary Zone Electrophoresis. J. Chromatogr, A, 1997; 773:311.

195. Bunke, A., Jira, T. Enantiomeric Separation by Capillary Electrophoresis Using Chiral and Achiral Ion Pairing Reagents. Pharmazic, 1996; 51:479.

196. Kuhn, R., Wagner, J., Walbroehl, Y., Bereuter, T. Potential and Limitations of an Optically Active Crown Ether for Chiral Separation in Capillary Zone Electrophoresis. Electrophoresis, 1994; 15:828.

197. Kuhn, R., Erni, F, Bereuter, T. Chiral Recognition and Enantiomeric Resolution Based on Host-Guest Complexation with Crown Ethers in Capillary Zone Electrophoresis. Anal. Chem., 1992; 64:2815.

198. Huang, W. X., Xu, H., Fazio, S. D., Vivilecchia, R. V. Chiral Separation of Primary Amino Compounds Using a Non-chiral Crown Ether with y8-Cyclodextrin in Capillary Electrophoresis. J. Chromatogr, B: Biomed. Appl, 1997; 695:157.

199. Armstrong, D. W, Chang, L. W, Chang, S. S. C. Mechanism of Capillary Electrophoresis Enantioseparations Using a Combination of an Achiral Crown Ether Plus Cyclodextrins. J. Chromatogr, A, 1998; 793:115.

200. Armstrong, D. W, Rundlett, K. L., Chen, J.-R. Evaluation of the MacrocycUc Antibiotic Vancomycin as a Chiral Selector for Capillary Electrophoresis. Chirality, 1994; 6:496.

201. Gasper, M. P, Berthod, A., Nair, U. B., Armstrong, D. W Comparison and Modeling Study of Vancomycin, Ristocetin A, and Teicoplanin for CE Enantioseparations. Anal. Chem., 1996; 68:2501.

202. Trelh-Siefert, L. A., Risley D. S. Capillary Electrophoretic Enantiomeric Separations of Nonsteroidal Anti-inflammatory Compounds Using the Macrocychc Antibiotic Actaplanin A and 2-Methoxyethanol. J. Liq. Chromatogr Related Technol, 1998; 21:299.

203. Reilly, J., Risley, D. S. The Separation of Enantiomers by Countercurrent Capillary Electrophoresis Using the MacrocycUc Antibiotic A82846B. LC/GC, 1998; 16:170.

204. Hong, M., Soini, H., Baker, A., Novotny M. V Complexation between Amylodextrin Oligomers and Selected Pharmaceuticals Measured through Capillary Electrophoresis. Anal. Chem., 1998; 70:3590.

205. Fillet, M., Hubert, P, Crommen, J. Enantioseparation of Non-steroidal Anti-inflammatory Drugs by Capillary Electrophoresis Using Mixtures of Anionic and Uncharged p-Cydodex-trins as Chiral Additives. Electrophoresis, 1997; 18:1013.

206. Nishi, H., Nakamura, K., Nakai, H., Sato, T. Enantiomeric Separation of Drugs by Mucopolysaccharide-Mediated Electrokinetic Chromatography. Anal. Chem., 1995; 67:2334.

207. Cole, R. O., Sepaniak, M. J., Hinze, W L. Optimization of Binaphthyl Enantiomer Separations by Capillary Zone Electrophoresis Using Mobile Phases Containing Bile Salts and Organic Solvent. HRC & CC, 1990; 13:579.

References 2 0 7

208. Aumatell, A., Wells, R. J. Enantiomeric Differentiation of a Wide Range of Pharmacologically Active Substances by Cyclodextrin-Modified Micellar Electrokinetic Capillary Chromatography Using a Bile SaU. J. Chromatogr., A, 1994; 688:329.

209. Iran, A. D., Blanc, T., Leopold, E. J. Free Solution Capillary Electrophoresis and Micellar Electrokinetic Resolution of Amino Acid Enantiomers and Peptide Isomers with L- and D-Marfey's Reagents. J. Chromatogr., 1990; 516:241.

210. Chan, K. C , Muschik, G. M., Issaq, H. J. Enantiomeric Separation of Amino Acids Using Micellar Electrokinetic Chromatography after Pre-column Derivatization with the Chiral Reagent l-(9-Fluorenyl)-ethyl chloroformate. Electrophoresis, 1995; 16:504.

211. Lurie, I. S. Micellar Electrokinetic Capillary Chromatography of the Enantiomers of Amphetamine, Methamphetamine and Their Hydroxyphenethylamine Precursors. J. Chromatogr, 1992; 605:269.

212. Anhoff, M., Einarsson, S., Chiral Deriyatization, in Chiral Liquid Chromatography, Chapter 4, Ed. W J. Lough, 1990, Blackie & Sons: p. 39.

213. Guzman, N. A. Column Watch. On-Line Bioaffinity, Molecular Recognition, and Preconcen-tration in CE Technology LC/GC, 1999; 17:16.

214. Chu, Y.-H., Kirby, D. P, Karger, B. L. Free Solution Identification of Candidate Peptides from Combinational Libraries by Affinity Capillary Electrophoresis/Mass Spectrometry. J. Am. Chem. Soc, 1995; 117:5419.

215. Gomez, E A., Avila, L. Z., Chu, Y.-H., Whitesides, G. M. Determination of Binding Constants of Ligands to Proteins by Affinity Capillary Electrophoresis: Compensation for Electroosmotic Flow. Anal. Chem., 1994; 66:1785.

216. Dalluge, J. J., Sander, L. C. Precolumn Affinity Capillary Electrophoresis for the Identification of Clinically Relevant Proteins in Human Serum: Application to Human Cardiac Troponin 1. Anal. Chem., 1998; 70:5339.

217. Akashi, M., Sawa, T., Baba, Y, Tsuhaku, M. Specific Separation of Oligodeoxynucleotides by Capillary Affinity Gel Electrophoresis (CAGE) Using Poly(9-vinyladenine)-Polyacrylamide Conjugated Gel. J. High Res. Chromatogr, 1992; 15:625.

218. Alber, J. K., Reddy K. R., Lee, C. S. Post-capillary Affinity Detection of Protein Microhetero-geneity in Capillary Zone Electrophoresis. J. Chromatogr, A, 1997; 759:139.

219. Busch, M. H. A., Carels, L. B., Boelens, H. E M., Kraak, J. C , Poppe, H. Comparison of Five Methods for the Study of Drug-Protein Binding in Affinity Capillary Electrophoresis. J, Chromatogr, A, 1997; 777:311.

220. Busch, M. H. A., Kraak, J.C, Poppe, H. Principles and Limitations of Methods Available for the Determination of Binding Constants with Affinity Capillary Electrophoresis. J. Chromatogr, A, 1997; 777:329.

221. Okun, V. M., Bilitewski, U. Analysis of the Biotin-Binding Protein Actinavidin Using Affinity Capillary Electrophoresis. Electrophoresis, 1996; 17:1627.

222. Amini, A., Westerlund, D. Evaluation of Association Constants Between Drug Enantiomers and Human aj-Acid Glycoprotein by Applying a Partial-Filling Technique in Affinity Capillary Electrophoresis. Anal. Chem., 1998; 70:1425.

223. Heegaard, N. H. H., Mortensen, H. D., Roepstorff, P Demonstration of a Heparin-Binding Site in Serum Amyloid P Component Using Affinity Capillary Electrophoresis as Adjunct Technique. J. Chromatogr, A, 1995; 717:83.

224. Gunnarsson, K., Valtcheva, L., Hjerten, S. Capillary Zone Electrophoresis for the Study of the Binding of Antithrombin to Low-Affinity Heparin. GlycoconjugateJ., 1997; 14:859.

225. Barker, G. E., Russo, P, Hartwick, R.A. Chiral Separation of Leucovorin with Bovine Serum Albumin Using Affinity Capillary Electrophoresis. Anal. Chem., 1992; 64:3024.

226. Avila, L. Z., Chu, Y. H., Blossey E. C , Whitesides, G. M. Use of Affinity Capillary Electrophoresis to Determine Kinetic and Equilibrium Constants for Binding of Arylsulfonamides to Bovine Carbonic Anhydrase. J. Med. Chem., 1993; 36:126.


227. Shimura, K., Kasai, K.-i. Determination of the Affinity Constants of Concanavalin A for Monosaccharides by Fluorescence Affinity Probe Capillary Electrophoresis. Anal. Biochem., 1995; 227:186.

228. VanderNoot, V. A., Hileman, R. E., Dordick, J. S., Linhardt, R. J. Affinity Capillary Electrophoresis Employing Immobilized Glycosoaminoglycan to Resolve Heparin-Binding Peptides. Electrophoresis, 1998; 19:437.

229. German, I., Buchanan, D. D., Kennedy, R. T. Aptamers as Ligands in Affinity Probe Capillary Electrophoresis. Anal. Chem., 1998; 70:4540.

230. Ahmed, A., Ibrahim, H., Pastore, E, Lloyd, D. K. Relationship Between Retention and Effective Selector Concentration in Affinity Capillary Electrophoresis and High-Performance Liquid Chromatography. Anal. Chem., 1996; 68:3270.

231. Schmitt, P, Freitag, D., Trapp, I., Garrison, A. W, Schiavon, M., Kettrup, A. Binding of s-Tri-azines to Dissolved Humic Substances: Electrophoretic Approaches Using Affinity Capillary Electrophoresis (ACE) and Micellar Electrokinetic Chromatography (MEKC). Chemosphere, 1997; 35:55.

232. Lausch, R., Reif, O.-W, Riechel, P, Schleper, T. Analysis of Immunoglobulin G Using a Capillary Electrophoretic Affinity Assay with Protein A and Laser-Induced Fluorescence Detection. Electrophoresis, 1995; 16:636.

233. Heegaard, N. H. H. Determination of Antigen-Antibody Affinity by Immunocapillary Electrophoresis. J. Chromatogr, A, 1994; 680:405.

234. Baba, Y., Inoue, H., Tsuhako, M., Sawa, T., Kishida, A., Akashi, M. Evaluation of the Selective Binding Ability of Oligodeoxynucleotides to Poly(9-vinyladenine) Using Capillary Affinity Gel Electrophoresis. Anal. Sci., 1994; 10:967.

235. Chu, Y. H., Whitesides, G. M. Affinity Capillary Electrophoresis Can Simultaneously Measure Binding Constants of Multiple Peptides to Vancomycin. J. Org. Chem., 1992; 57:3524.

236. Dunayevskiy Y. M., Lyubarskaya, Y. V, Chu, Y.-H., Vouros, P, Karger, B. L. Simultaneous Measurement of Nineteen Binding Constants of Peptides to Vancomycin Using Affinity Capillary Electrophoresis-Mass Spectrometry. J. Med. Chem., 1998; 41:1201.

CHAPTER

Capillary Isoelectric Focusing

5.1 5.2 5.3 5.4 5.5 5.6 5.7 5.8 5.9 5.10 5.11 5.12 5.13

Basic Concepts Separation Mechanism pH Gradient Formation Electrode Buffer Solutions Resolving Power Capillaries and Reagents Performing a Run Injection Focusing Mobilization Salt Effects Detection Applications References

5.1 BASIC CONCEPTS

A fundamental parameter of electrophoresis is mobility. This is defined by the charge-to-mass ratio of a solute. Should a solute become neutral during the run, migration will cease. During the course of a run, modification of mobility can occur in a pH gradient generated online by a series of reagents known as carrier ampholytes. Therein lies the fundamental premise of isoelectric focusing (lEF).

Conventional lEF is performed in an anticonvective medium such as the slab gel. The advantages and disadvantages of gels, discussed in Section 1.1, apply equally to lEE In contrast to the usual gel format, the pore size of an lEF gel should be sufficiently large to reduce the impact of molecular sieving, which otherwise would lengthen the run time. Agarose gels have an extremely large pore size, 500 nm for a 0.16% gel (1). Polyacrylamide gels with large pore sizes can be formulated as well.

209

210 Chapter 5 Capillary Isoelectric Focusing

Detection in the slab gel is time consuming and semiquantitative. The carrier ampholytes must be washed out of the gel to avoid reaction with the staining reagent. Small peptides are not detectable, since they are lost during the wash step (2). Gels are unnecessary in capillary isoelectric focusing (CIEF), since the ampholyte separation media is effectively supported and contained by the capillary walls. CIEF is performed in free solution.

Mobilization represents an additional difference between capillary and conventional lEF. Since the focusing process produces electrically neutral solutes, some means of eluting the bands is required for detection. In the slab gel, mobilization is unnecessary, since detection is performed by staining. Hjerten and Zhu's original work (3) describes a scheme that forces the proteins to move past the detector window. Such a technique is known as mobilization.


Figure 5.1 illustrates the mechanism of CIEF A sample is mixed with a series of reagents known as carrier ampholytes. Characteristic of ampholytes is good buffering capacity at their individual p/ values to "carry" the pH (4). In other words, the pK^ of each ampholyte is very close to its pi.

The capillary is filled with a sample-ampholyte blend, and a voltage is applied. Ampholyte concentrations of 0.5-2.0% are used for most applications. Under the influence of the applied electric field, carrier ampholytes have the capacity to generate a pH gradient along the length of the capillary Without considering at this stage how the gradient is formed, let us examine the behavior of a zwitterionic solute toward the pH gradient.

There are two possible behaviors. At a pH below the solute's pi, the zwitterion is positively charged and migrates toward the cathode. As the solute migrates through the pH gradient, it encounters progressively higher pH values. At some point, the solute enters a region along the gradient where the ampholyte pH is equal to its own pi. At this point, the solute's net charge becomes zero, and migration ceases.

CATHODE / ^ ANODE

'^^^^^^^^^^^^^M^M

^^^^^^^^^^^^^^^^^S.

pH GRADIENT HIGH pH " LOW pH

FIGURE 5.1 Illustration of the capillary isoelectric focusing process.


The second possible behavior occurs when the solute is negatively charged. This occurs along the pH gradient when the pH is greater than the solute's pi. The solute migrates toward the anode, encountering progressively lower pH values. While the direction of migration through the pH gradient is opposite to the case just described, the net result is the same. Solute migration ceases whatever the direction of approach to the proteins pi. When a capillary is filled with a mixture of ampholytes and solutes, both behaviors occur, as shown in Figure 5.1. CIEF is a true focusing technique. If a solute diffuses into a buffer region where it becomes charged, it will migrate back to the region of zero charge.

The CIEF system is substantially different from zone electrophoresis. In CZE, the buffer system is homogeneous throughout the length of the capillary and throughout the duration of the run. In CIEF, a heterogeneous pH gradient is created inside the capillary by applying voltage across the carrier ampholytes. The breadth of the pH gradient depends on which series of ampholytes is selected.

Ampholytes are commercially available to cover both wide and narrow pH ranges, as shown in Figure 5.2.i The Ampholines (LKB-Pharmacia) are a series of polyamino-polycarboxylic acids. Servalytes (Serva) are polyamine-polysul-fonic acids. Pharmalytes (LKB-Pharmacia) are branched polyamino-polycarboxylic acids.

The mechanism of isoelectric focusing permits the separation of solutes based on their isoelectric points. Most reported applications are for proteins,

1 Carrier ampholytes are available from Bio-Rad (Richmond, CA), Pharmacia LKB (Piscataway, NJ), Serva Biochemicals (Paramus, NJ), Sigma Chemical (St. Louis, MO), and Beckman Instruments (Fullerton, CA).

pH 2 3 4 5 6 7 8 9 1 0 1 1

FIGURE 5.2 Carrier ampholyte pH ranges. Each horizontal hne represents the pH range covered by an ampholyte blend.


but any zwitterionic solute can be separated via this approach. Using CIEF, there is no need to develop a buffer system to separate solutes based on mobility. If the solutes have sufficiently different pJ values, they will separate. This mechanism of separation contrasts sharply with CZE and CGE, where the bases for separation are respectively the charge-to-mass ratio and molecular size. Righetti's elegant textbook (1) is recommended for further background on conventional lEF methodology Many of the phenomena observed in the slab gel are directly related to occurrences in the capillary

5.3 PH GRADIENT FORMATION

Before applying the voltage, the individual ampholytes are uniformly distributed throughout the capillary The pH is uniform as well and represents the average pH of the ampholyte blend (Figure 5.3). Individual ampholytes may be cationic or anionic at the start of the run, depending on their pi values. This is illustrated in Figure 5.4 (top), where charge is assigned to the individual ampholytes based on a starting pH of 7.

When voltage is applied, the ampholyte mixture begins to separate into individual components (Figure 5.4, bottom). Complete separation of adjacent ampholytes should not be attained, since this would cause a discontinuity in the pH gradient. Either a step gradient would be observed, or in extreme cases, if the zones separate completely, a water zone would occur. At this point, the electric field strength becomes so high that the capillary sustains damage.

pH

pH GRADIENT

AMPHOLYTE CONCENTRATIONS

DISTANCE ALONG CAPILLARY FIGURE 5.3 Representation of a pH gradient and individual ampholyte concentrations before focusing.

5.4 Electrode Buffer Solutions 213

CATHODE ANODE

iOH-*-i a-^g* s-^s* 4-"*'io* 7 6-1 •H+| I ; *• •• •• _ • ! !

HIGHliH AVERAGE pH IS 7 L O W H

CATHODE ANODE

J0H-*.| 10 9 8 7 6 5 4 3 | •H*

HIGH pH LOW ~pH

FIGURE 5.4 The process of pH gradient formation.

Positively charged (high-pl) ampholytes migrate toward the negative electrode, and negatively charged ampholytes toward the positive electrode. The migration of ampholytes causes the electrolyte pH to increase as the cathode is approached. Conversely, the pH in the anodic region begins to decline as the low-pI (negatively charged) ampholytes migrate in that direction.

Eventually, migration will cease as each ampholyte encounters a pH where its net charge becomes zero. This occurs at the individual isoelectric point, or pi, of each ampholyte. If a solute such as a protein is present in the carrier ampholyte blend, it too migrates while charged, until it encounters a pH equivalent to its pi. The system does not distinguish between a protein and a carrier ampholyte; both behave as zwitterions and migrate according to their respective pi values. At steady state, the ampholyte distribution and resulting pH gradient are illustrated in Figure 5.5. The measured current should approach but never reach zero, since no further movement from either ampholytes or solutes is expected, except diffusion.

5.4 ELECTRODE BUFFER SOLUTIONS

The pH values of the respective electrode buffer solutions are critical in CIEE The anodic buffer, or anolyte, must have a pH that is lower than the pi of the most acidic ampholyte. Similarly, the cathodic buffer, or catholyte, must have a higher pH than the most basic ampholyte. If these conditions are not met, ampholytes will bleed into an electrode buffer reservoir. Diluted versions of electrode solutions used in conventional lEF are appropriate for CIEE Selection of the proper anolyte and catholyte serves as a barrier to prevent the migration of


pH

pH GRADIENT

DISTANCE ALONG CAPILLARY FIGURE 5.5 Representation of a pH gradient and individual ampholyte concentrations after focusing.

ampholytes into the reservoirs. Should an acidic ampholyte migrate into the more acidic anolyte, its charge becomes positive, and it migrates back toward the negative electrode. The corresponding process occurs at the negative electrode. It is important that fresh sodium hydroxide be used. Otherwise, uptake of carbon dioxide may lower the pH, causing a cathodic drift of the pH gradient (5).

Sodium hydroxide (20 mM) and phosphoric acid (10 mM) solutions are frequently used as catholyte and anolyte solutions, particularly with the pH 3-10 gradient. The 2:1 ratio of hydroxideiacid is not coincidental. Studies have shown that this provides for the most stable pH gradient. Otherwise, anodic or cathodic drifts in the pH gradient are more likely to occur (4). The mechanism for drift is based on a phenomenon known as isotachophorsis (ITP). These drifts reflect the loss of ampholytes into the electrode reservoirs. The 2:1 ratio provides for symmetrical drift of the gradient. For narrow-range gradients such as pH 6-8, weaker acids and bases such as 40 mM glutamic acid (anolyte) and 40 mM arginine (catholyte) can be used (6). Avoiding alkaline pH prolongs the lifetime of the coated capillary

While not studied by CIEF, ampholyte blends that are more acidic or basic than the carrier ampholytes have been used as the anolyte and/or catholyte in slab-gel lEF (1). The same holds true for Good's buffer solutions of the appropriate pi. HEPES (pJ 7.51) is used as the anolyte for narrow-range alkaline gradients (1).

5.5 RESOLVING POWER

The resolving power, Apl, of CIEF is described by (1)

5.6 Capillaries and Reagents 2 1 5

DCdpH/^)^ (5.1)

^E(dAt/dpH)

where D is the diffusion coefficient, E is the field strength, ^ is the mobihty of the protein, and d/J^/d pH describes the mobihty-pH relationship. The term d pH/dx represents the change in the buffer pH per unit of capillary length. This adjustable parameter is controlled by selecting an appropriate ampholyte pH range as well as the capillary length. For highest resolution, a narrow-pH ampholyte range should be selected. To separate a wide pi range of proteins, although at lower resolution, select a broad-pH-range ampholyte blend. Under optimal conditions, resolution of 0.02 pH units can be achieved (1).

5.6 CAPILLARIES AND REAGENTS

A. COATED CAPILLARIES

The use of coated capillaries is preferred in CIEF, to suppress the EOF and minimize protein adsorption (3, 7, 8). The reduction of EOF is desired in CIEF for several reasons:

1. The EOF may sweep the solutes past the detector before focusing is complete. This is another form of cathodic drift of the pH gradient.

2. Silanol ionization at the capillary wall is pH dependent. In the presence of a pH gradient, the degree of ionization will vary along with the pH. As a result, the measured EOF is the average value derived from the integrated contributions along the entire length of the capillary. This can generate an osmotic pressure throughout the capillary, causing a hydro-dynamic-like flow profile. However, the focusing effect from CIEF probably overcomes this form of band broadening.

3. In the presence of EOF; the low-pH anolyte enters the capillary, since the net flow is directed toward the cathode. As more of the anolyte fills the capillary, the EOF declines as ionized silanol groups are protonated. This leads to nonlinearity of the pH gradient. Under nonlinear conditions, the migration time is not proportional to the pi. Calibration is difficult when this occurs, since multiple standards are required. In addition, the migration time for acidic proteins becomes rather long.

A wide variety of coated capillaries can be employed in CIEF This is illustrated in Figure 5.6, where six different capillaries were investigated for the separation of recombinant tissue plasminogen activator (rt-PA) (9). All of these capillaries except the DB WAX were considered equivalent. The success of these


o

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FIGURE 5.6 Effect of the capillary coating type on the resolution of rt-PA glycoforms. Capillary: 27 cm (20 cm to detector) x 50 |lm id coated as specified; ampholytes: 1.5% pH 3-10,1.5% pH 5-8, 0.1% HPMC, 0.75% TEMED, 4 M urea; anolyte: 10 mM phosphoric acid; catholyte: 20 mM sodium hydroxide; focusing: -500 V/cm; mobilization: one-step; detection: UV, 280 nm; temperature: 20°C. Reprinted with permission from J. Chromatogr, 717, 61 (1995), copyright © 1995, Elsevier Science Publishers.

capillaries may be due to the presence of urea and hydroxypropylmethylcellu-lose (HPMC) in the ampholyte blend. The use of urea may eliminate solubility problems that the protein may have in and around its isoelectric point.

When selecting a capillary, it is important to measure its stability, since many of the coatings are susceptible to degradation by alkaline reagents. It is usually necessary to change the capillary after 50-100 runs. Shortening the degree of contact with alkaline reagents is very effective in reducing this problem. If the

5.6 Capillaries and Reagents 217

protein has a low pJ, then ehminating the alkaUne ampholytes via a narrow-range gradient is even more effective.

Since the migration times may change with time and can be influenced by the solute concentration and salt concentration, the use of internal standards to calibrate the pH gradient is mandatory.

B. INTERNAL STANDARDS

Marker proteins or synthetic markers are generally employed to calibrate pi versus time. Kits are available from many manufacturers. Typical lEF protein markers are listed in Table 5.1. A good marker protein provides a single or several well-defined sharp peaks. Crude protein preparations often give broad bands due to impurities or microheterogeneity. The problem with proteins as internal standards is interference with the solutes of interest in complex separations. When this occurs, it is necessary to perform two runs: with and without the internal standards.

Small zwitterionic molecules such as azo dyes are alternative choices. For example, methyl red (pi 3.8) does not absorb at 280 nm, the protein-monitoring wavelength. Using a diode array detector, it is possible to monitor simultaneously the methyl red absorbance at 500 nm and the protein absorbance at 280 nm. In this regard, methyl red is an ideal internal standard, but few such molecules have been identified.

Aminomethylphenyl dyes absorb at 280 nm, although their maximum absorption is around 400 nm (10). Nevertheless, they have been employed as

Table 5.1 Marker Proteins for lEF

Protein

Amyloglucosidase

Glucose oxidase

Trypsin inhibitor

p-Lactoglobulin A

Carbonic anhydrase II

Carbonic anhydrase II

Carbonic anhydrase I

Myoglobin

Lentil lectin

Trypsinogen

Source

Aspergillus niger

Aspergillus niger

Soybean

Bovine milk

Bovine erythrocytes

Bovine erythrocytes

Human erythrocytes

Horse heart

Lens culinaris

Bovine pancreas

Pl

3.6

4.2

4.6

5.1

5.4

5.9

6.6

6.8, 7.2

8.2, 8.6, 8.8

9.3

Data excerpted from Sigma Chemical catalog 1998, p. 1985.


pi markers in CIEF (11). The advantage of synthetic markers is that their pi values are not affected by the presence of denaturants such as urea. In complex protein separations, the use of 400-nm detection allows calibration of the system even if the peaks are obscured at 280 nm.

C. PREPARATION OF METHYLCELLULOSE SOLUTION

To further suppress the EOF and maintain a coated capillary wall, methylcellulose solutions are useful for preparing the anolyte and ampholyte solutions. If a partial-fill technique is employed (Section 5.8), the catholyte can be prepared in methylcellulose as well. The viscous media also helps suppress peak distortions during the subsequent mobilization steps. The following procedure works well for solubilizing this material (12). Alternatively, hydroxypropylmethylcel-lulose (HPMC) solutions can be used. For one-step CIEF, a 0.1% HPMC solution lowers the EOF without completely suppressing it. This material is also more water soluble than methylcellulose.

Heat 250 mM HPCE-grade water to 60-70°C. Add 0.9375 g (0.375%) methylcellulose (1500 cp for a 2% aqueous solution at 20°C, Sigma) and mix to form a suspension. After 5 min, place the suspension in ice-water and stir until the material enters solution and the mixture reaches 5-10°C. It is essential to filter this material through a 0.45-|Lim filter prior to use.

D. AMPHOLYTE BACKGROUND U V ABSORPTION

Unfortunately carrier ampholytes contain some chemicals that absorb at 280 nm. These reagents were not originally designed for capillary electrophoresis. The ampholytes were created for the slab gel with detection by traditional staining methods using dyes such as Coomassie brilliant blue R-250. When running in the capillary format, it is essential to perform a blank run.

For hydrophilic proteins with good absorbance at 280 nm, the ampholyte backgrounds are usually not a problem, since one can start with a solute concentration of several hundred micrograms per milliliter. If the proteins are hydrophobic, then the protein concentration must be reduced to avoid precipitation. If the protein is deficient in aromatic amino acids and absorbs poorly at 280 nm, interference from the ampholytes, even at 0.5% concentration, may be a problem.

Figure 5.7 shows CIEF of rt-PA versus the blank for several ampholytes from various manufacturers (13). Substantial backgrounds are found for some of the reagents. Since successful separations were reported elsewhere for both Bio-Lytes and Servalytes, it is possible that batch-to-batch variation is occurring.


Time (minutts)

FIGURE 5.7 Impact of carrier ampholytes on the blank (dashed line) and the rt-PA glycoform profile. Capillary: 27 cm (20 cm to detector) x 50 |im id eCAP neutral; ampholytes: 1.5% pH 3-10, 1.5% pH 5-8, 0.1% HPMC, 0.75% TEMED, 4 M urea; anolyte: 10 mM phosphoric acid; catholyte: 20 mM. sodium hydroxide; focusing: -500 V/cm; mobilization: one-step; detection: UV, 280 nm; temperature: 20°C; rt-PA concentration: 125 jLig/mL. Key: A) Pharmalytes, B) Ampholines, C) Bio-Lytes, D) Servalytes. Reprinted with permission from J. Chromatogr., 744, 279 (1996), copyright © 1996, Elsevier Science Publishers.

When ampholyte interference is a problem, decolorizing activated carbon can be used to clean the ampholytes (14). Stop up a Pasteur pipet with a plug of glass wool and fill with some activated carbon. Add the ampholytes and, using a pipet bulb, push the material through the column. Hopefully the aromatic ampholytes will adsorb on the activated carbon, thereby reducing the reagent background.

All ampholyte blends are not created equal. If ampholytes from one source give a high background, try the same pH range from another manufacturer. Beware that since ampholytes are not tested for CIEF, batch-to-batch variation in the reagent background may occur when different lots, even from the same manufacturer, are employed.


E. PREPARATION OF NARROW-RANGE

AMPHOLYTE MIXTURES

Narrow-range pH gradients are advantageous for two reasons:

1. If the pH range is less basic, the capillary coating is better preserved. 2. The resolution is improved.

The improvement in resolution is shown in Figure 5.8 for the separation of rt-PA using a blend of pH 4-8 and pH 3-10 ampholytes (9). The optimal blend is a 50:50 mixture of the two ampholyte solutions.

The need for blended ampholyte solutions when running narrow-range gradients arises from the number of carrier ampholytes present. When the blends are manufactured, the narrow-range cuts contain fewer ampholyte species. While there are at least hundreds of ampholytes present per pH unit (1), the reduction in the total number of ampholyte species has consequences. This can lead to the production of step gradients or water zones within the capillary. The use of the ampholyte blend ensures an adequate number of ampholytes. The higher concentration of the narrow-range material serves to improve the resolution.

If more resolution is required at a particular pH range, it is possible to flatten the gradient at that point through the addition of a spacer or separator (15). For example, the separation of adult hemoglobin (Hb A) from its glycated form, Hb Ale, is difficult, because the ApJ is 0.03 pH units. The addition of 330 mM /3-alanine and 330 mM 6-aminocarproic acid to the narrow-range ampholyte blend (5% 6-8, 0.5% 3-10) provides a baseline separation (16). The trick is to

MIGRATION TIME rMIN^

FIGURE 5.8 Effect of changing the ratio of ampholytes from 100% pH 5-8 to 100% pH 3-10. Other conditions as per Figure 5.7. Reprinted with permission from J. Chromatogr, 717, 61 (1995), copyright © 1995, Elsevier Science Publishers.


add a separator with a pJ nearly equal to the portion of the gradient to be flattened. The reason for the high concentrations of spacers is that they are poor ampholytes. That is, their pK^ values are far from their pi values. But there are no available good ampholytes that precisely hit the pH 6.9 region. Good ampholytes function well as spacers at the 15 mM level. Other good ampholytes that can serve as spacers include aspartic acid, pi 2.77; glutamic acid, pi 3.22; triglycine, pi 5.59; histidine, pi 7.47; and lysine, pi 9.74. Refer to reference (15) for a complete listing of ampholytes that can serve as spacers.

F. ADDITIVES FOR HYDROPHOBIC PROTEINS

The tendency of hydrophobic proteins to aggregate and precipitate is a major problem in lEF, whether in the slab-gel or capillary format. The focusing power of CIEF produces an increase in solute concentration by a factor of more than 200 (17). Proteins also readily precipitate as the pi is approached, since their charge and, thus, their electrostatic repulsion approach zero. The problem is exacerbated by the necessity of increasing protein concentration to visualize minor impurities (15).

Protein precipitation is indicated first by spikes in the electropherogram, followed by clogging of the capillary Additives are required to suppress the aggregation of hydrophobic proteins to keep them in solution. Two excellent review articles describing solutions to this problem serve as the source for much of the material in the following discussion (18, 19).

Urea has been a reagent of first choice, but it is not without problems. Urea can suppress protein aggregation by disrupting hydrogen bonds, the so-called "iceberg effect." At high concentrations, urea is not uniformly dispersed, but forms organized channels. These channels bind to linear alkyl chains, but not branched or cyclic molecules. The complexes, which are undefined, are actually less soluble than the native protein. To better solubilize these solutes, surfactants are often used in combination with urea. Unfortunately, the high ionic strength of SDS may cause heating problems in CIEF. Nonionic surfactants such as Brij-35, Triton X-100, Nonidet P-40, octyl glucoside, and lauryl maltoside or zwitterionic detergents such as sulfobetains (CHAPS, for example) can be used at concentrations of 0.1-5%. Not all of these surfactants have been used in the capillary format. The surfactant must not absorb at 280 nm, which necessitates the use of the reduced form of Triton X-100.

Another important problem linked to the use of urea is carbamylation. In aqueous solution, urea is in equilibrium with ammonium cyanate, the level of which increases with increasing concentration and temperature. Cyanate can react with amines to produce substituted ureas. The use of ultrapure and fresh


urea and the use of low temperatures (20-25°C) minimizes the cyanate concentration. The ampholytes themselves serve as good cyanate scavengers.

Most of the reported work on CIEF has been on native proteins (we think), since the effects of solubilizers such as surfactants and urea have not been thoroughly studied. Denaturing CIEF has been reported (20) wherein 5% mercap-toethanol and 8 M urea were added to the ampholyte blend. SDS was not used in the separation, and it is not known whether the sample was boiled prior to CIEE The state of denaturation of these and other proteins is not well known.

Gentle detergents and other additives are more often used in lEF, and so it becomes more likely that native proteins are being separated. Polyols such as ethylene glycol, glycerol, sorbitol, or nonreducing sugars may be useful. A mixture of 20% sorbitol and 100 mM taurine in pH 6-8 ampholytes has been successful in separating thermamylase (21). Glycols do not keep thermamylase in solution, but 20% sucrose in taurine buffer works well. Thus, a strategy of mixing polyols and zwitterions proved successful in dealing with solubility problems.

5.7 PERFORMING A RUN

There are three basic modes of operation of CIEF, which are distinguished by the process by which the focused bands are mobilized through the detector. These are:

1. Chemical (salt) mobilization 2. Electroosmotic (one-step) mobilization 3. Hydrodynamic mobilization

The conditions for four separate protocols are summarized in Table 5.2. The mode of injection also serves to distinguish these methods. The capillary can be completely filled with sample-ampholyte blend or partially filled. In addition, the sample can be injected sandwiched between segments of ampholytes, thereby avoiding the need to dissolve the sample in ampholytes (2).

A. CAPILLARY CONDITIONING

Various rinsing procedures can be employed to ensure a clean and reproducible capillary surface. To ensure that residual proteins are removed from the capillary, a postrun wash consisting of a 1-min high-pressure rinse with 100 mM phosphoric acid followed by 0.5 min with methanol and 0.5 min with deion-ized water is employed (22). An alternative procedure is to rinse for 1 min with 0.1 N hydrochloric acid, 1 min with 10 mM phosphoric acid, and 1 min with water (9).

3.7 Perform

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B. SAMPLE PREPARATION

Dissolve the protein in 0.5-2% carrier ampholytes to give a final concentration of between 50 and 500 |ig/mL. Always start at the lower concentrations to minimize the potential of precipitation upon focusing. Hydrophilic proteins can be run at the higher concentrations, hydrophobic proteins at lower concentrations. Note: It is useful to have a syringe handy to unclog the capillary.

If the salt content of a protein sample is above 50 mM, the sample should be desalted or online desalting should be utilized (Section 5.11). One such desalting method is as follows (23):

1. Add 5 parts cold methanol to 1 part sample. 2. Store at -80°C for 30 min. 3. Centrifuge at 15,000g. 4. Discard the supernatant. 5. Resuspend the sample pellet in the ampholyte solution.

Dialysis, gel filtration, dilution, or solid-phase extraction are other common desalting techniques. These procedures are covered in Section 10.5.

5.8 INJECTION

Since viscous liquids are being loaded, much time will be saved if the instrument can be operated in a high-pressure mode. At 200 psi, a 50-|Lim-i.d. capillary can be filled with 0.4% methylcellulose solution in 30 s.

A. WHOLE-CAPILLARY INJECTION

The entire capillary can be filled with sample-ampholyte blend. It is possible that material will focus on the blind side of the capillary beyond the detector window. In this case, add N,N,N,N-tetramethylenediamine (TEMED) to the ampholyte blend to block the blind side of the capillary This basic reagent is positively charged at most pH values, and so it migrates toward and occupies the space near the end of the capillary (6, 24). The concentration of TEMED can be as high as 7.5% (9), though 0.5-2% is most common. It is important to verify with markers that you are getting the expected pJ range.

B. PARTIAL-CAPILLARY INJECTION

The following protocol is designed for the Beckman P/ACE CE system (25) and must be adapted for other instruments as described subsequently

5.9 Focusing 225

1. Fill the capillary (eCAP neutral), 27 cm (20 cm to detector) x 50 |iL with catholyte (20 mM sodium hydroxide in 0.4% methylcellulose) for 0.4 min at 20 psi.

2. Load the sample-ampholyte blend for 0.4 min at 20 psi. The length of the ampholyte section in the capillary is about 16 cm, or 80% of the capillary length to detector.

It is simple to accurately calibrate the length of time it takes to fill the capillary with the sample-ampholyte blend. By following the following procedure, the method can be adapted to any instrument:

1. Set the detector to 200 nm. 2. Set the operating temperature of the system (usually 20-25°C). 3. Flush the capillary with water, anolyte or catholyte depending on protocol. 4. Inject sample-ampholyte blend, and note the time the detector signal

increases. Decrease the measured time by 10-20%, and use that for the load time.

C. PLUG INJECTION

Another approach permits injection of proteins dissolved in water. This procedure is utilized for the no longer manufactured ABl 270A (2), but it can be adapted to any instrument. The pressures and timing are set for a 72 cm (50 cm to detector) x 50 jiim i.d. capillary.

1. Fill the capillary with catholyte (20 mM sodium hydroxide in 0.4% methylcellulose) for 8 min with 20" vacuum.

2. Load the ampholyte solution for 4.5 min with 20" vacuum. 3. Inject a sample and marker separately, each for 30 s with 5" vacuum. 4. Inject a plug of ampholyte solution for 6 s at 20" vacuum to insulate the

sample.

At this point, the leading edge of the ampholyte blend is near but not past the detector. The advantage of this method is the ability to inject aqueous samples, thereby sparing material that would otherwise have to be dissolved in ampholytes.

The sensitivity of the method is reduced compared with the other injection techniques, since very little protein (20 nL) is injected into the capillary. Using whole-capillary injection with a 20 cm x 50 |im i.d. capillary, the injection size is 400 nL.

5.9 FOCUSING

The next step in CIEF is focusing (see Figure 5.1). When the voltage is first activated, the current is relatively high, since both ampholytes and solutes are

2 2 6 Chapter 5 Capillary Isoelectric Focusing

highly mobile at the start of the run. As the focusing proceeds, the pH gradient forms, along with solute migration to the appropriate position along the gradient. When the ampholytes and solutes approach their respective pi values, their mobilities begin to slow, and as a result, the current declines.

Focusing takes only a 2-5 min at 400-600 V/cm. Monitoring the current is a useful indicator for optimizing the focusing time. Typically, the current will decline to and level off at a few microamperes. Overfocusing results in protein precipitation, as shown by spikes on the electropherogram (24, 26). Underfo-cusing can result in split peaks for a single protein (19).

Overfocusing can also cause damage to the capillary as a result of the formation of water zones. These zones tend to occur near the cathode and are more prevalent when acidic gradients are run. Repeated capillary fractures were observed when a field strength of 1200 V/cm was employed for 5 min. Reducing the field strength to 600 V/cm resolved the problem without substantial changes in speed and resolution.

Most runs are performed in the positive-polarity mode, where the more basic proteins elute first. To switch the order of elution, operate in the negative-polarity mode. The catholyte is placed on the inlet side and the anolyte on the outlet side of the capillary This is opposite to what is conventionally done. Under these conditions, the more acidic proteins elute first. This mode of operation is particularly important when one-step and chemical mobilization are used, since the acidic side of the capillary is not effectively mobilized.

5.10 MOBILIZATION

With focusing complete, it is necessary to mobilize or elute the contents of the capillary past the detector to record the electropherogram. There are three ways to accomplish this:

1. Electrophoretic (salt) mobilization 2. Hydrodynamic mobilization 3. Electroosmotic (one-step) mobilization

Methods 1 and 2 isolate focusing and mobilization into separate and discrete processes. For method 1, it is necessary to turn off the voltage when switching buffer reservoirs. In method 3, focusing and mobilization occur simultaneously.

A. ELECTROPHORETIC (SALT) MOBILIZATION

This form of mobilization uses salts (3) or zwitterions (27) as additives to a buffer reservoir to effect pH changes in the capillary when the voltage is applied. The direction of mobilization can be anodic or cathodic, as indicated in Figures 5.9 and 5.10, depending on which reservoir the salt is added to. Cathodic mobi-

5.10 Mobilization 227

DETECTOR WINDOW

CATHODE \ ANODE

I PROTESNS AND AMPHOLYTES •

NaOH OH"

NaCI

FIGURE 5.9 Anodic mobilization.

lization is usually selected, unless very acidic proteins at the far end of the capillary need to be seen. As mobilization proceeds, the current always increases.

Electrophoretic mobilization occurs by adding salt to one of the buffer reservoirs (27). Since electroneutrality is required, this condition is satisfied by (5)

C„. + IC. NH3+ C^u- + ZrfC^ (5.2)

where CH% CQH-, CNH3% ^^^ ^coo' ^^e the concentrations in equivalents per liter of all charged species in the ampholyte-solute blend.

If a salt (anion) is added to the catholyte, the expression becomes

Cj + + 2C j^ „^ = Q H - + ^^-COO" + C (5.3)

where C^^- represents an anion (m is the charge on that anion). The added salt, now the anion, competes with OH" for electromigration into the capillary. Since fewer hydroxyls enter the capillary, the pH declines. Proteins previously at their pi values become cationic and begin to migrate toward the cathode.

For anodic mobilization, an expression equivalent to Eq. (5.3) is

C^n^ + C^+ H- 2Cj^^^+ - CQ^- -H Z C , "COO" (5.4)

where Cx»* represents the cation added to the anolyte. The added anion is of no consequence, since it never migrates into the capillary. The same holds true for

CATHODE /

DETECTOR WINDOW

ANODE

^^sSlEiSi^

I PROTEINS AND AMPHOLYTES i |

NaOH NaC!

OH - *

cr—I •H^ H3PO4

FIGURE 5.10 Cathodic mobilization


the cation when using cathodic mobihzation. Increasing the concentration of salt speeds the mobihzation process at the expense of thermal deformation due to Joule heating.

Examples of cathodic and anodic mobilization for human hemoglobin and transferrin are shown in Figure 5.11 (3). This early work employed 3-mm-i.d. capillaries that were rotated at 40 rpm to reduce convection. While the applied field strength is low and speed of separation relatively slow by today's standards, the fundamental aspects of CIEF are well illustrated. As expected, the electro-pherograms are approximate mirror images of each other. Better resolution is always seen during the early portions of the electropherograms.

The relationship of the change in pH versus the distance from the end of a slab gel is shown in Figure 5.12 (27) for anodic mobilization. The pH changes are substantial along the first 4 cm of the gel and then converge to no measurable differences. This means the relationship between the migration time and pJ is not Unear. The loss of linearity is a consequence of mobilization, not of focusing.

The failure to effectively mobilize solutes at the far end of the capillary is addressed in part with a zwitterionic mobilization reagent. When cathodic mobilization is employed, the problem occurs with acidic (low-pl) proteins. Adding to the catholyte a zwitterion (instead of salt) with a pi lower than the most acidic protein is effective in mobilizing the far end of the capillary (24). During mobilization, the low-p7 zwitterion migrates near the anode until it becomes neutral. Ampholytes and solutes with greater pi values all acquire a positive charge and mobilize toward the cathode.

Zwitterionic mobilizers can eliminate unwanted peaks from the extremes of electropherograms. For example, performing cathodic mobilization with pi 6.9

Hb Tr

(min.) 10

FIGURE 5.11 Chemical (salt) mobilization of human hemoglobin and transferrin by capillary CIEE Capillary: agarose-plugged, methylcellulose-coated 38 cm x 3 mm i.d. rotated at 40 rpm; ampholytes: 1% solution of Pharmalyte (pH 3-10); anolyte: phosphoric acid, 20 mM; catholyte: sodium hydroxide, 20 mM; focusing: 1200 V for 20 min; mobilization: (a) anolyte: 20 mM sodium hydroxide; catholyte: 20 mM sodium hydroxide; (b) anolyte I: 20 mM phosphoric acid; catholyte: II: 20 mM phosphoric acid; mobilization voltage: 3000 V; detection: Uy 280 nm. Reprinted with permission from J, Chromatogr., 346, 265 (1985), copyright © 1985, Elsevier Science Publishers.

5.10 Mobilization 229

10

PH

2 4 8

DISTANCE (cm)

FIGURE 5.12 The pH gradient at different mobilization times on a 2-mm slab gel (3%C, 6%T) cast in a 2.5% solution of Bio-Lyte (pH 3-10) on a water-cooled microscope slide. The pH measurements were made with a surface electrode. Focusing: with 20 mM sodium hydroxide as catholyte and 20 mM phosphoric acid as anolyte; mobilization: anodic with 20 mM phosphoric acid with 80 mM sodium chloride as the anolyte. Reprinted with permission from J. Chromatogr., 387, 127 (1987), copyright © 1987, Elsevier Science Publishers.

zwitterion eliminates all bands with pi values below 6.9 (24). For anodic mobilization, all bands with pi values above that of the zwitterion will not be detected.

B. HYDRODYNAMIC MOBILIZATION

The problem of mobilization of proteins at the far end of the capillary is addressed by hydrodynamic mobilization (3). After focusing, the capillary contents are eluted using an HPLC pump. Hydrodynamic band broadening is reduced by applying the focusing voltage during elution.

Chen and Wiktorowicz (2) and later Huang et al. (25) adapted this approach with modern instrumentation. Instead of pumping out the capillary contents, high-precision vacuum or pressure is used. The focusing voltage is applied during the mobilization step and need not be turned off as the pressure or vacuum is applied. A linear relationship between mobility and pi occurs over the full pH range. The technique works best when low pressure (0.5 psi, 50 mbs) or vacuum (5" Hg) is used.

C. ELECTROOSMOTIC MOBILIZATION

In this technique, EOF mobilization occurs together with the focusing step (6, 26, 28-30). The EOF is reduced by adding 0.1% methylcellulose to the ampholyte blend. This is sufficient to ensure that focusing occurs before any solute migrates past the detection window. The advantage here is that buffers

2 3 0 Chapter 5 Capillary Isoelectric Focusing

do not need changing nor does the voltage have to be turned off and on. While bare silica capillaries have been employed here, it is better to use a coated capillary to produce a more linear pH gradient. The electropherograms shown in Figures 5.6-5.8 were generated using this technique.

D. COMPARISON OF THE MOBILIZATION

TECHNIQUES

Figure 5.13 compares the three mobilization techniques for a series of standard proteins (20). The highest resolution is found for chemical mobilization at the expense of run time. The opposite is true for single-step CIEF Hydrodynamic mobilization represents the middle ground between the two other techniques and is simple to implement. Both chemical and hydrodynamic mobilization produce linear pH gradients versus time. This is not the case for single-step CIEF The reproducibility of all three techniques is good, particularly when internal standards are used to compensate for capillary coating degradation and the variable salt content of the sample (20).

E. BUFFER DEPLETION

Since buffers are not generally added to the anolyte and catholyte, pH changes can occur during the course of repeated runs. This problem is solved by frequently changing these solutions. The use of a zwitterionic catholyte such as taurine, pH 8.8, also reduces this problem since the reagent has buffering capacity, but this Umits the operating range of the system to proteins with pJ values below 8.5 (31).

5.11 SALT EFFECTS

In Figure 5.4, the formation of a pH gradient in the absence of salt is illustrated. The impact of added salt is now shown in Figure 5.14. The presence of salt ions is significant, since these ions are quite mobile and thus carry much of the current in the capillary. In this regard, they are competitive with the carrier ampholytes and delay the onset gradient formation. As Figure 5.14 (bottom) illustrates, a compression of the gradient occurs. Basic proteins exhibit longer migration times, whereas acidic proteins show shorter times.

While desalting of samples is often necessary, the use of a voltage ramp permits the high-mobility salt ions to exit the capillary prior to the formation of the pH gradient (32, 33). A voltage gradient of 5-10 kV applied over 6 min, followed by focusing at 15-20 kV over 5-10 min, is effective to desalt protein samples

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FIGURE 5.13 Comparison of mobilization techniques. Chemical mobilization (top)—capillary: jLlSIL DB-1, 27 cm (20 cm to detector); 4% Pharmalyte 3-10, 1% Temed, 0.8% MC; anolyte: 20 mM phosphoric acid; catholyte: 20 mM sodium hydroxide (focusing) + 30 mM sodium chloride (mobilization); focusing: 5 min at 20 kV; cathodic mobililization at 20 kV. Hydrodynamic mohilization (middle)—capillary: [iSlL DB-I, 27 cm (20 cm to detector); ampholytes: 4% Pharmalyte 3-10, 1% Temed, 0.8% MC; anolyte: 10 mM phosphoric acid, 0.4% MC; catholyte: 20 mM sodium hydroxide; focusing: 10 min at 10 kV; mobilization: 0.5 psi at 10 kV One-step CIEF (bottom)—capillary: eCAP neutral 37 cm (30 cm to detector); ampholytes: 4% Pharmalyte 3-10, 1.5% Temed, 0.4% HPMC; anolyte, 10 mM phosphoric acid; catholyte 20 mM sodium hydroxide; voltage: -10 kV Key: (1) cytochrome c; (2) ribonuclease; (3) myoglobin; (4) carbonic anhydrase; (5) /^-lactoglobulin. The dotted line is the current. Reprinted with permission from Electrophoresis, 16, 2121 (1995), copyright © 1995 Wiley-VCH.

232

CATHODE

Chapter 5 Capillary Isoelectric Focusing

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containing 200 mM sodium chloride. The vohage should be increased at such a rate to prevent the current from exceeding 15 |LlA, or resolution degrades. As Figure 5.15 shows, the resolution appears to improve despite the presence of 200 mM salt (32).

5.12 DETECTION

A. UV DETECTION

Carrier ampholytes absorb UV light below 250 nm. For most modes of HPCE, the UV background is not a big problem, since the optical path length is short and the buffer is homogeneous. While a high-UV reagent background increases the noise of the system, the baseline can be zeroed.

In CIEF, the electrolyte composition is continually changing during focusing and subsequent mobilization. Monitoring the electropherogram at 200 nm produces a substantial background, masking all but the most concentrated proteins in the sample (26).

This problem precludes the use of 200 nm for detection, the most sensitive wavelength for proteins. Sufficient selectivity toward proteins is obtained at 280 nm. The loss of sensitivity at this wavelength compared with that at 200 nm depends on the aromatic amino acid composition of the protein. Proteins with few aromatic residues yield poor sensitivity. Limits of detection usually fall into the low micrograms per milliliter range. This is often limited by the reagent background signal from the carrier ampholytes themselves.

5.12 Detection 233

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FIGURE 5.15 Online desalting using a voltage gradient, (a) Absence of salt; (b) sample in 200 mM sodium chloride. Capillary: 37 cm (30 cm to detector) x 50 |Xm i.d., PVA-coated (homemade); ampholytes: 2% 3-10; anolyte: 50:49:1 (methanol:water:acetic acid); catholyte: 50:49:1 (methanol:water:ammonium hydroxide); focusing: 20 kV for 10 min; mobilization anolyte: catholyte solution; detection: 280 nm. Reprinted with permission from Anal. Chem., 69, 2786 (1997), copyright © 1997 Am. Chem. Soc.


The use of the diode-array detector (DAD) in conjunction with coated capillaries may reduce the capillary lifetime. Since the DAD passes all wavelengths of light through the capillary, the coating may be damaged. An interference filter that removes all but the 280-nm wavelength band should solve this problem.

B. LASER-INDUCED FLUORESCENCE

Laser-induced fluorescence (LIF) has been utilized in CIEF both for native fluorescence of hemoglobins in single red blood cells (34) and for tetramethyl-rhodamine-derivatized anti-human growth hormone antibody (35). Limits of detection by LIF are in the sub-ng/mL range.

C. MASS SPECTROMETRY

The combination of CIEF and mass spectrometry is analogous to 2-D electrophoresis (36). In this case, the mass spectrometer provides the molecular weight information instead of SDS-PAGE. This information can be obtained by online CIEF-MS (37) or by using CIEF as a micropreparative technique (38, 39).

For the online system, CIEF is performed conventionally in a 20-cm capillary mounted inside an electrospray probe. After focusing, the outlet reservoir (catholyte) is removed and the capillary tip set to 0.5 mm outside of the probe. A sheath liquid of 50% methanol, 49% water, and 1% acetic acid (pH 2.6) pumped with a syringe pump at 3 |xL/min produces a stable electrospray. Cathodic mobilization is produced by changing the inlet (anolyte) from 20 mM phosphoric acid to the sheath liquid. During mobilization, two power supplies are used, one for the capillary (10 kV) and another for the electrospray (5 kV).

The ampholyte ions were observed up to m/z 800 and thus did not interfere with the protein signals. Increasing the ampholyte concentration caused a reduction in the protein ion counts, and this is protein dependent. Good results are found using 0.5% ampholytes.

D. WHOLE-CAPILLARY IMAGING

An instrument designed to image the entire capillary is available from Convergent Bioscience Ltd, Etobicoke, Ontario, Canada. The instrument employs UV detection with imaging provided by a charge-coupled device. Since the entire capillary is continuously monitored, the mobilization step is no longer required (40-42).

5.13 APPLICATIONS

Table 5.3 lists a variety of applications, buffer recipes, and operating conditions. The rest of this section will examine a few applications in more detail.

5.13 A

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A. HEMOGLOBINS

Perhaps the most widespread apphcation of lEF in the slab gel is hemoglobin analysis. There are hundreds of variations of human hemoglobin that result from single-point amino acid mutations. Abnormal hemoglobin is found in 1 of 10,000 individuals with electrophoresis as the diagnostic test. CIEF is useful for performing a high-speed separation isolating some important variants such as hemoglobin S, the sickle cell variant that results from a 6 Glu —> Val replacement. Zhu et al. (24) resolve hemoglobins S, C, F, and A in 6 min using salt mobilization (Figure 5.16). The separation of hemoglobin A from F is remarkable, since the two proteins differ by only 0.05 pi units.

The work of Zhu was improved on by Hempe et al. (22, 49) in several significant ways. The combination of hydrodynamic mobilization and detection at 415 nm greatly simplified the experiments. Detection at 415 nm instead of 280 nm removes any potential interferences from carrier ampholytes or other endogenous proteins in blood serum. The use of a narrow-range pH 6-8 gradient improves resolution as well. Results were reported for a wide variety of hemoglobin mutations, a few of which are reproduced in Figure 5.17.

A (7 JO)

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FIGURE 5.16 Separation of hemoglobin variants by CIEF in a 12 cm x 25 |Lim i.d. coated capillary using pH 3-10 ampholytes. Focusing and mobilization were carried out at 8 kV. Protein concentration: 250 |lg/mL of each protein; detection: 280 nm; isoelectric points are: hemoglobin A, pi 7.1; F, pi 7.15; S, pi 7.25; and C, pi, 7.5. Reprinted with permission from J. Chromatogr., 559, 479 (1991), copyright © 1991 Elsevier Science Publishers.

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FIGURE 5.17 Separation of hemoglobin variants by CIEF with detection at 415 nm. Capillary: JLISIL DB-1 coated 27 cm (20 cm to detector) x 50 \im i.d.; ampholytes: Pharmalyte 2% of 6-8:3-10 (10:1) in 0.375% MC; anolyte: 100 mM phosphoric acid in 0.375% MC; catholyte: 20 mM sodium hydroxide; injection: 10-30 s \ov^ pressure (<50 nL); focusing: 5 min at 30 kV; mobilization: lov^ pressure with 30 kV voltage. Courtesy of James Hempe, LSU School of Medicine.


B. RiBONUCLEASES

Chen and Wiktorowicz (2) separated, in part, RNase T (pj 2.9), RNase ab (pi 9.0), and site-directed mutants (pJ 3.1, 3.1, 3.3) of RNase T using the hydro-dynamic mobihzation method. The two mutants of pi 3.1 were not separable. If there are any differences between these two, a narrow-range acidic gradient may be required for separation.

The separation, shown in Figure 5.18, includes the marker proteins RNase A (pi 9.5), carbonic anhydrase (pi 5.9), j8-lactoglobulin (pi 5.1), and CCK-Flank-ing protein (pi 2.75). The authors reported the DB-1 capillary was stable for at least 420 runs over a two-month period. The combination of the DB-1 capillary and the methylcellulose additive may account in part for this stability. If the coating is harmed by base, the methylcellulose polymer may serve to recoat the damaged section.

When using marker proteins for internal standardization, the pi measurement precision ranged from 0.5% to 3.3% RSD. A peak area calibration curve was linear versus injection time (volume) from 10 to 80 s (50 to 400 nL).

Mobilixation Time iwcdn)

FIGURE 5.18 Capillary CIEF of RNase ab, RNase T , and RNase T site-directed mutants. Capillary: DB-1 coated 72 cm (50 cm to detector) X 50 |im i.d.; ampholytes: Servalyte 3-10, 0.5% with 0.4% methylcellulose; injection: 30 s vacuum (200 nL); focusing: 6 min at 30 kV; mobilization: vacuum (5" Hg) with 30 kV voltage; detection: UV, 280 nm; solutes: 200 |ag/mL each (4 ng injected). Reprinted with permission from Anal Biochem., 206, 84 (1992), copyright © 1992 Academic Press.


C. RT-PA METHOD VALIDATION

Moorhouse et al. (46), from Genentech, investigated the validation of a CIEF method for rt-PA. Separations have aheady been shown in Figures 5.6-5.8, and so no others will be given here. The analytical conditions employed are described in the caption of Figure 5.6.

This method separates the glycoforms of rt-PA based on their sialic acid content. Nine to 10 peaks are consistently resolved, with the profile dependent on the class of ampholyte used. The pJ of the glycoforms ranged from 6.5 to 7.5.

Method validation involves a series of controlled experiments designed to measure a particular attribute of the method, such as accuracy, specificity, precision, and ruggedness (see Section 10.7). Only those methods meeting acceptable criteria are deemed suitable to submit to regulatory authorities and to use in quality control to assess purity, potency, and identity.

The pH gradient was characterized with synthetic markers that bracketed the pH range of rt-PA. Since one-step mobilization was used, the pH gradient was not linear with time, particularly for the acidic portion of the gradient. The migration times of the synthetic markers were affected by the presence of rt-PA, which means that calculated pi depends on the nuances of the particular system. Ideal markers would not be affected by the protein.

Peak area linearity was examined from 25 to 1000 |ig/mL and found to be acceptable, with a correlation coefficient of 0.997. The limit of detection was 50 |ig/mL, which corresponds to 25 ng of protein. The limit of detection using Coomassie brilliant blue stain was 2 |Lig via the slab gel. Recovery was assessed by fraction collection, with the collected fractions measured by an ELISA method. A 110% recovery was found, which indicates no material was retained on the capillary. Run-to-run migration time precision ranged from 2.3% to 3.1%. Peak area precision ranged from 0.6% to 10.4%, with only the first two peaks giving RSDs of greater than 3%. Day-to-day peak migration time precision was 6-8%, and peak area precision ranged from 1% to 14%, with peak 1 being poor once more. Migration times decreased with each passing day, which was a function of the deterioration of the coating. Since the EOF provides for mobilization, the use of hydrodynamic mobilization might remedy this problem. It is also possible that the DB-1 coating might prove superior.

Ruggedness was assessed by examining five different lots of capillaries and three capillaries from the same lot. Two lots of capillaries showed prolonged run times. This is a consequence once again of one-step mobilization, since it depends on the EOF Increasing the voltage from 400 to 600 V/cm gave shorter migration times but essentially the same pattern. Varying the temperature from 15°C to 30°C also affected the migration times but not the patterns. Increasing the TEMED concentration from 0.38% to 1.5% lengthened the run but did not alter the pattern. Pharmalytes and ampholines gave different patterns by CIEF but also did the same on the slab gel.

References 241

Since the only troublesome criteria was the day-to-day precision, the authors of this work are considering using hydrodynamic mobiUzation (with complete EOF suppression) so as not to be hampered by changes in the EOF of the system. Recently, a validated method for identification of a monoclonal antibody has been implemented at Genentech (50).

REFERENCES

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4. Liu, X., Sosic, Z., Krull, I. S. Capillary Isoelectric Focusing as a Tool in the Examination of Antibodies, Peptides and Proteins of Pharmaceutical Interest. J. Chromatogr, A, 1996; 735:165.

5.Hjerten, S. Isoelectric Focusing in Capillaries, in Capillary Electrophoresis Theory and Practice, Eds. P. D. Grossman and J. C. Colburn. 1992, Academic Press. 191.

6.Mazzeo, J. R., Krull, 1. S. Capillary Isoelectric Focusing of Proteins in Uncoated Fused-Silica Capillaries Using Polymeric Additives. Anal. Chem., 1991; 63:2852.

T.Hjerten, S., Elenbring, K., Kilar, E, Liao, J. L., Chen, A.J. C , Siebert, C. J., Zhu, M. D. Carrier-Free Zone Electrophoresis, Displacement Electrophoresis and Isoelectric Focusing in a High-Performance Electrophoresis Apparatus. J. Chromatogr, 1987; 403:47.

8.Bolger, C. A., Zhu, M., Rodriguez, R., Wehr, T. Performance of Uncoated and Coated Capillaries in Free Zone Electrophoresis and Isoelectric Focusing of Proteins. J. Liq. Chromatogr, 1991; 14:895.

9.Moorhouse, K. G., Eusebio, C. A., Hunt, G., Chen, A. B. Rapid One-Step Capillary Isoelectric Focusing Method to Monitor Charged Glycoforms of Recombinant Human Tissue-Type Plasminogen Activator. J. Chromatogr, A, 1995; 717:61.

lO.Caslavska, J., Molteni, S., Chmelik, J., Slais, K., Matulik, E, Thormann, W. Behaviour of Substituted Aminomethylphenol Dyes in Capillary Isoelectric Focusing with Electroosmotic Zone Displacement. J. Chromatogr, A, 1994; 680:549.

11. Kilar, E, Vegvari, A., Mod, A. New Set-up for Capillary Isoelectric Focusing in Uncoated Capillaries. J. Chromatogr, A, 1998; 813:349.

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13. Chen, A. B., Rickel, C. A., Flanigan, A. H. G., Moorhouse, K. G. Comparison of Ampholytes Used for Slab Gel and Capillary Isoelectric Focusing of Recombinant Tissue-Type Plasminogen Activator Glycoforms. J. Chromatogr, A, 1996; 744:279.

14. Pritchett, T. Personal communication. 1998. 15.Righetti, P G., Bossi, A., Gelfi, C. Capillary Isoelectric Focusing and Isoelectric Buffers: An

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Colony Stimulating Hormone by Capillary Electrophoresis, Capillary Isoelectric Focusing and High-Performance Liquid Chromatography. J. Chromatogr, 1993; 652:215.

18. Rabilloud, T. Solubilization of Proteins for Electrophoretic Analysis. Electrophoresis, 1996; 17:813.


19.Rodriguez-Diaz, R., Wehr, T., Zhu, N. Capillary Isoelectric Focusing. Electrophoresis, 1997; 18:2134.

20.Schwer, C. Capillary Isoelectric Focusing: A Routine Method for Protein Analysis. Electrophoresis, 1995; 16:2121.

21. Conti, M., Galassi, M., Bossi, A., Righetti, R G. Capillary Isoelectric Focusing: The Problem of Protein SolubiUty J. Chromatogr, A, 1997; 757:237.

22.Hempe, J. M., Granger, J. N., Warrier, R. R, Graver, R. D. Analysis of Hemoglobin Variants by Capillary Isoelectric Focusing. J. Capillary Electrophor, 1997; 4:131.

23.Reif, O. W, Freitag, R. Control of the Cultivation Process of Antithrombin III and Its Characterization by Capillary Electrophoresis. J. Chromatogr, A, 1994; 680:383.

24. Zhu, M., Rodriguez, R., Wehr, T. Optimizing Separation Parameters in Capillary Isoelectric Focusing. J. Chromatogr, 1991; 559:479.

25. Huang, T.-L., Shieh, P C. H., Cooke, N. Isoelectric Focusing of Proteins in Capillary Electrophoresis with Pressure-Driven Mobilization. Chromatographia, 1994; 39:543.

26.Thormann, W, Caslavska, J., Molteni, S., Chmelik, J. Capillary Isoelectric Focusing with Elec-troosmotic Zone Displacement and On-Column Multichannel Detection. J. Chromatogr, 1992; 589:321.

27.Hjerten, S., Liao, J.-L., Yao, K. Theoretical and Experimental Study of High-Performance Elec-trophoretic Mobilization of Isoelectrically Focused Protein Zones. J. Chromatogr, 1987; 387:127.

28.Mazzeo, J. R., Krull, I. S. Examination of Variables Affecting the Performance of Isoelectric Focusing in Uncoated Capillaries. J. Microcol. Sep., 1992; 4:29.

29.Mazzeo, J. R., Krull, I. S. Improvements in the Method Developed for Performing Isoelectric Focusing in Uncoated Capillaries. J. Chromatogr, 1992; 606:291.

30.Mazzeo, J. R., Martineau, J. A., Krull, I. S. Performance of Isoelectric Focusing in Uncoated and Commercially Available Coated Capillaries. Methods, 1992; 4:205.

31.Rodriguez-Diez, R., Zhu, M., Wehr, T. Strategies to Improve Performance of Capillary Isoelectric Focusing. J. Chromatogr, A, 1997; 772:145.

32. Clarke, N. J., TomUnson, A. J., Schomburg, G., Naylor, S. Capillary Isoelectric Focusing of Physiologically Derived Proteins with Online Desalting of Isotonic Salt Concentrations. Anal. Chem., 1997; 69:2786.

33. Clarke, N. J., Tomlinson, A. J., Naylor, S. Online Desalting of Physiologically Derived Fluids in Conjunction with Capillary Isoelectric Focusing-Mass Spectrometry J. Am. Soc. Mass Spectrom., 1997; 8:743.

34.Lillard, S. J., Yeung, E. S. Analysis of Single Erythrocytes by Injection-Based Capillary Isoelectric Focusing with Laser-Induced Native Fluorescence Detection. J. Chromatogr, B: Biomed. Appl, 1996; 687:363.

35. Shimura, K., Kasai, K.-i. Fluorescence-Labeled Peptides as Isoelectric Point (pi) Markers in Capillary Isoelectric Focusing with Fluorescence Detection. Electrophoresis, 1995; 16:1479.

36. Tang, Q., Harrata, A. K., Lee, C. S. Two-Dimensional Analysis of Recombinant E. Coh Proteins Using Capillary Isoelectric Focusing Electrospray Ionization Mass Spectrometry. Anal Chem., 1997; 69:3177.

37. Tang, Q., Kamel Harrata, A., Lee, C. S. Capillary Isoelectric Focusing-Electrospray Mass Spectrometry for Protein Analysis. Anal. Chem., 1995; 67:3515.

38. Grimm, R. Micropreparative Capillary Isoelectric Focusing of Protein and Peptide Samples Followed by Protein Sequencing. J. Capillary Electrophor, 1995; 2:111.

39.Foret, F, Muller, O., Thorne, J., Gotzinger, W, Karger, B. L. Analysis of Protein Fractions by Micropreparative Capillary Isoelectric Focusing and Matrix-Assisted Laser Desorption Time-of-Flight Mass Spectrometry. J. Chromatogr, A, 1995; 716:157.

40. Wu, J., Pawliszyn, J. Application of Capillary Isoelectric Focusing with Adsorption Imaging Detection to the Quantitative Determination of Human Hemoglobin Variants. Electrophoresis, 1995; 16:670.

References 243

41. Wu, J., Pawliszyn, J. Protein Analysis by Isoelectric Focusing in a Capillary with an Absorption Imaging Detector. J. Chromatogr., B: Biomed. Appl, 1995; 669:39.

42. Palm, A., Lindh, C , Hjerten, S., Pawliszyn, J. Capillary Zone Electrophoresis in Agarose Gels Using Absorption Imaging Detection. Electrophoresis, 1996; 17:766.

43.Schnabel, U., Fischer, C.-H., Kenndler, E. Characterization of Colloidal Gold Nanoparticles According to Size by Capillary Electrophoresis. J. Microcolumn Sep., 1997; 9:529.

44.Kundu, S., Fenters, C. Isoelectric Focusing of Monoclonal Antibodies by Capillary Electrophoresis. J. Capillary Electrophor, 1995; 2:273.

45.Thorne, J. M., Goetzinger, W. K., Chen, A. B., Moorhouse, K. G., Karger, B. L. Examination of Capillary Zone Electrophoresis, Capillary Isoelectric Focusing and Sodium Dodecyl Sulfate Capillary Electrophoresis for the Analysis of Recombinant Tissue Plasminogen Activator. J. Chromatogr, A, 1996; 744:155.

46. Moorhouse, K. G., Rickel, C. A., Chen, A. B. Electrophoretic Separation of Recombinant Tissue-Type Plasminogen Activator Glycoforms: Validation Issues for Capillary Isoelectric Focusing Methods. Electrophoresis, 1996; 17:423.

47.Kilar, F, Hjerten, S. Separation of the Human Transferrin Isoforms by Carrier Free High-Per-formance Zone Electrophoresis and Isoelectric Focusing. J. Chromatogr, 1989; 480:351.

48. Schmitt, P, Poigner, T, Simon, R., Freitag, D., Kettrup, A., Garrison, A. W. Simultaneous Determination of Ionization Constants and Isoelectric Points of 12 Hydroxy-s-Triazines by Capillary Zone Electrophoresis and Capillary Isoelectric Focusing. Anal. Chem., 1997; 69:2559.

49.Hempe, J. M., Granger, J. N., Craver, R. D. Capillary Isoelectric Focusing of Hemoglobin Variants in the Pediatric Clinical Laboratory. Electrophoresis, 1997; 18:1785.

50. Hunt, G., Hotaling, T, Chen, A. B. Validation of a Capillary Isoelectric Focusing Method for the Recombinant Monoclonal Antibody C2B8. J. Chromatogr, A, 1998; 800:355.

CHAPTER 6 Size Separations in Capillary Gels and Polymer Networks

6.1 Introduction 6.2 Separation Mechanism 6.3 Materials for Size Separations 6.4 Size Separations with Nonreplaceable Polyacrylamide 6.5 Size Separations with Replaceable Agarose 6.6 Introduction to Polymer Networks 6.7 Operating Characteristics of Polymer Networks 6.8 Additional Materials for Polymer Networks 6.9 Detection 6.10 Operating Hints Using Polymer Networks 6.11 Applications and Methods Development 6.12 Reducing the Problem of Biased Reptation

References

6.1 INTRODUCTION

Slab-gel electrophoresis is the predominant technique for the separation of peptides, proteins, and polynucleotides. The slab-gel format provides mechanical stability for the separation, reduces solute dispersion from convection and diffusion, and permits handling for detection, scanning, storage, and so forth, as described in Section 1.1. Unlike lEF gels, w^hich exist solely for these functions, slab gels also provide the mechanism for separation.

Gels are porous matrices comprising polymeric materials dissolved in a solvent, usually an aqueous buffer. The pore size of the gel is determined by the concentration of the polymeric reagent and the three-dimensional structure of the matrix. Chemical crosslinkers further influence structural rigidity and pore size of the gel w^hen incorporated during the polymerization process. The porous structure of the gel provides effective separations of macromolecules based on molecular size.

245

246 Chapter 6 Size Separations in Capillary Gels and Polymer Networks

True gels are composed of lyophilic (solvent-loving) colloidal particles, the best known of which is polyacrylamide. Lyophobic (solvent-hating) colloidal materials such as agarose are known as sols. Agarose solutions must be prepared in boiling aqueous buffer to dissolve sufficient material to form the gel matrix.

The transition from the slab-gel to the capillary format began in 1983 (1) when Hjerten filled a 150-|im-i.d. tube with polyacrylamide. Later, the first reports of SDS-PAGE appeared (2, 3). Though many applications have been reported using rigid crosslinked gels, usually polyacrylamide, the separations were not robust owing to repeated gel failure.

In slab-gel electrophoresis, a gel is poured and polymerized just before use.^ The gel must be sufficiently viscous to eliminate flow and permit handling. Tackiness of the material must also be avoided. The gel is generally designed to be used once in the slab-gel format. The field strength is kept relatively low to minimize problems with heat dissipation, even for ultrathin and cooled gels.

In the capillary format, high-viscosity gels are prepared by crosslinking them in situ (4--6) or by pumping into the capillary under high pressure (7). The crosslinked gels must be stable enough to tolerate multiple runs, as the entire capillary must be replaced if the gel deteriorates. Unfortunately, the high applied field strength used in HPCE can cause abrupt and unpredictable failures. Generation of air bubbles and/or cracking of the gel matrix results in the loss of electrical continuity and the termination of the separation.

It became clear that a pumpable matrix is required for the capillary format. The advantage of these replaceable gels is that a fresh matrix can be employed for each run. This required the development of high-pressure pumping systems to blow the material into the capillary. For example, the PE Biosystems Prism 310 employs a high-pressure syringe to pump the viscous reagent at pressures as high as 1800 psi. All of the commercial capillary-array DNA sequencers now use replaceable polymer networks.

It was recognized early on that noncrosslinked polymeric materials such as methylcellulose derivatives, polyethylene glycols, linear polyacrylamide, and dextrans can also define molecular pores. These systems are best described as polymer networks, entangled polymers, or physical gels. Despite the physical differences between rigid crosslinked gels and polymer networks, the mechanisms of separation appear to quite similar.


The size separation mechanism is illustrated in Figure 6.1 for a series of oligonucleotides (or oligosaccharides). Driven by the electric field, solutes

iRehydratable gels are also commercially available, but these have no role in HPCE.


migrate toward the appropriate electrode through the gel (polymer) matrix. Small molecules pass through the pores unimpeded. Larger biopolymers may travel a tortuous path, moving through the pores in a snakelike fashion. Under properly controlled conditions, the solute's mobility is inversely proportional to its size.

Several mechanisms for the migration of macromolecules through polymer networks have been described (8). As shown in Figure 6.2, the Ogsten model treats a molecule as a nondeformable sphere, with the migration velocity determined by a solute's mobility modified by the probability of an encounter with a restricting pore. Solutes with a radius of gyration less than or equal to the average pore size (such as SDS-proteins) behave in this manner.

Large biopolymers do not necessarily follow the Ogston model. These molecules can deform during transit through the porous network. The movement of a long strand of DNA wriggling reptilelike through the polymer matrix is known as reptation. It is also observed that fragment resolution decreases with the size of the molecules. Large molecules such as DNA and oligosaccharides align with the electric field in a size-dependent manner that is biased toward the larger strands. The dependence of mobility on the molecular size is obscured by this process, which is known as biased reptation. This effect limits the size of DNA molecules that can be separated using conventional slab-gel techniques. Beyond 20,000 base pairs (bp), separations become difficult and pulsed-field electrophoretic techniques are usually employed (9, 10) This equipment is not presently available for commercial capillary electrophoresis instrumentation.

A further level of complexity in the mechanism of separation is the interaction of the macromolecule with the polymers used to form the porous network.

DIRECTION OF MIGRATION

FIGURE 6.1 Pictorial description of the mechanism of size separation by HPCE.


OGSTEN MODEL REPTATION

FIGURE 6.2 Comparison of the size separation mechanism for proteins (Ogsten model) and for DNA or oligosaccharides (reptation).

It has been shown that separations can occur at polymer concentrations far below what is required to define pores (11).

Gels and polymer networks separate the analyte based on molecular size. To correlate mobility to molecular weight, the macromolecule must be denatured to ensure that all solutes have the same charge-to-mass ratio. DNA and RNA are usually denatured by heating in formamide at 90°C for a few minutes. This improves both the separation and sizing accuracy.

For proteins, denaturation entails reduction of disulfide bonds and unfolding by heating with a solution composed of sodium dodecyl sulfate (SDS) and a reducing agent, /J-mercaptoethanol or dithiothreitol (DTT). After these processes, most proteins have roughly the same shape and charge density. SDS binds to and denatures proteins via electrostatic and hydrophobic interactions; about 1.4 g SDS is bound per gram of protein. Also, SDS is anionic, so that all proteins become negatively charged and migrate toward the anode. Under these conditions, a protein's mobility is proportional to its molecular weight, and the system can be calibrated with a sizing standard. SDS-PAGE is the standard method for determining size of proteins. When the molecular weight of a protein is less than 10 kDa, the SDS-binding stoichiometry may change, resulting in errors when calculating molecular weight (12).

6.3 MATERIALS FOR SIZE SEPARATIONS

A variety of reagents can be employed to develop a microporous network within the capillary. Classical reagents such as polyacrylamide and agarose have been adapted to the capillary format. Other low-viscosity solutions not suitable for slab gels, including linear polyacrylamide, polyethylene oxide, and methylcel-lulose, work well in capillaries. Both crosslinked and linear polymers can define

6.4 Size Separations with Nonreplaceable Polyacrylamide 249

pores in solution (Figure 6.3), either through covalent bonding or polymeric entanglement. These materials are classified as chemical and physical gels or polymer networks, respectively The rigid, crosslinked chemical gels are now only rarely used in HPCE.

When using low-viscosity materials, a coated capillary is frequently required to suppress the EOF. Otherwise, solutes may elute in a reverse order—that is, large molecules elute before small ones. Migration time imprecision and wall effects may also occur when using untreated capillaries. In other cases, the high viscosity of the polymer network suppresses the EOF, and a bare silica capillary can be used. This is usually the preferable choice.

The materials used for size separations can be considered either nonreplaceable or replaceable. It is clearly advantageous to replace the material in the capillary with each run. There are still a few capillaries containing nonreplaceable media that are commercially available.^

6.4 SIZE SEPARATIONS WITH NONREPLACEABLE POLYACRYLAMIDE

First introduced in 1959 by Raymond and Weintraub with later work by Orenstein and Davis, polyacrylamide has become a standard material for slab-gel electrophoresis (13). For capillary electrophoresis, the gel absorbs in the low-UV portion of the spectrum, resulting in detection problems when applied to proteins.

2J&W Scientific, Folsom, CA.

CHEMICAL GEL PHYSICAL GEL

FIGURE 6.3 Illustration of the differences between chemical and physical gels. Pores in chemical gels are defined via covalent bonds. In physical gels, polymeric entanglement defines the porous network.


The gel composition of this electrically neutral material is defined by %T, the total amount of acrylamide, and %C, the amount of crosslinker. Percent T is calculated by

o/T, acrylamide (g) + bisacrylamide (g)

lOOmL

Percent C is

o/oC = bisacrylamide(g) ^ ^^^ ^^^^

bisacrylamide (g) + acrylamide(g)

Bisacrylamide is frequently the crosslinker, though there are many alternatives. The pore size of the gel is controlled by both %T and %C. Highly crosslinked gels are usually employed to increase the pore size. A 30%C gel has a pore size of 200-250 A (14).

The preparation of a stable, bubble-free gel-filled capillary is a complex undertaking. Bubble production and gel shrinkage during polymerization are often confounding problems that do not occur in the open environment of the slab gel. The integrity of the polymerized gel often fails due to the high field strengths that are employed in HPCE. Gel failure can occur without warning, a significant problem during automated unattended runs. On the other hand, spectacular separations of oligonucleotides, such as that illustrated in Figure 6.4, have been reported. Table 6.1 lists some applications using rigid gels. However, it is anticipated that few users will opt for these materials.

6.5 SIZE SEPARATIONS WITH REPLACEABLE AGAROSE

Agarose is a complex group of polysaccharides extracted from the agarocytes of Rhodophyceae, a marine algae found predominately in the Pacific and Indian Oceans. Neutral, pyruvated, and sulfated fractions have been isolated, though all fractions contain some charged groups. The material is insoluble in cold water and slowly soluble in hot water. A 1% solution forms a stiff gel upon cooling. In 1961, Hjerten first employed agarose as a support for zone electrophoresis.

Righetti (14) summarized the properties of this remarkable material. The polysaccharide is a double helix with a pore structure more rigid than a poly-acrylamide strand. Even in dilute concentrations, the agarose structure has high mechanical strength. Pores with diameters of 500-800 nm have been reported. It is not surprising that agarose is useful in separating large segments of DNA. Even more remarkable is the low viscosity of this material; it is pumpable, a sig-

6.5 Size Separations with Replaceable Agarose 251

20 25 30 35 Ttfiie {mm)

FIGURE 6.4 CGE of poly((iA)2o and poly(dA)4o_6o on a 9%T linear polyacrylamide gel. Capillary: 60 cm (45 cm to detector) X 75 \im i.d.; field strength: 308 V/cm; current: 8.8 jiA; buffer: 100 mM Tris-borate, pH 8.3, 2 mM EDTA, 7 M urea; injection: electrokinetic, 10 kV for 0.5 s; detection: UV, 260 nm. Reprinted with permission from J. Chromatogr., 516, 33 (1990), copyright © 1990 Elsevier Science Publishers.

nificant advantage in HPCE. Among the advantages of agarose are UV transparency and lower toxicity than that of polyacrylamide.

Work still continues using agarose, though it is unlikely that the material will find widespread use due to the superior results found using polymer networks. In recent developments, low-melting, low-gelling agarose was used to separate DNA fragments in a pulsed-field system (23) and in a conventional system (24). A mixture of hydroxyethylcellulose and agarose has also been reported (25).

Table 6.1 Applications with Polyacrylamide Crosslinked Gels in CGE

Application Gel Composition Buffer

0.1 M Tris-borate, 2.5 mM EDTA, 7 M urea, pH 7.6

90 mM Tris, 90 mM boric acid, 2.5 mM EDTA, 1.3 mM Temed, pH8.3

0.1 M Tris, 0.25 M boric acid, 7 M urea, pH 7.6

50 mM Tris, 50 mM boric acid. 7 M urea

0.1 M Tris, 0.25 M borate. 2 mM EDTA, pH 8.48

89 mM Tris, 69 mM boric acid, 7 M urea, 2 mM EDTA, pH 8.6

0.1 M Tris, 0.1 M boric acid. 7 M urea, pH 8.8

0.1 M Tris, 0.25 M borate, 7 M urea, pH 7.5

90 mM Tris-phosphate, 0.1% SDS, 8 M urea, pH 8.61

Reference

15

16

17

18

19

20

21

22

2

DNA sequencing 3%T, 5%C

DNA sequencing 4%T, 5%C

Oligonucleotides 6%T, 5%C

Oligonucleotides 7.5%T, 3.3%C

Oligosaccharides 25.5%T, 3%C

Oligothymidylic acids 2.5%T, 3.3%C

Polyadenylates 5%T, 5%C

Polydeoxycytidines 6%T, 3%C

Proteins 10-12.5%T, 3.3%C

6.6 INTRODUCTION TO POLYMER NETWORKS

Methylcellulose (MC) and its derivatives are the prototypical polymer networks. Following the first reports by Hjerten et a\. (26) and Zhu et al. ill), much work with polymer networks has been accomplished using this material or its derivatives. Grossman and Soane (28) found that the pore size in these polymer networks depends on the polymer concentration and the radius of the mesh-forming polymer chain.

When the polymer concentration is low, the polymers are isolated from one another and exhibit no overlap (Figure 6.5). In this example, 0 is the polymer concentration, and 0* is the overlap or entanglement threshold. As the polymer concentration is increased, the chains begin to overlap. Finally, at yet higher polymer concentration, the entangled network is formed.

The overlap threshold can be evaluated experimentally by monitoring the viscosity of the polymeric solution. A plot of viscosity versus <P* becomes nonlinear at a concentration of 0.29% hydroxyethylcellulose (28), indicating the onset of polymer overlap. Pore sizes of 223-350 A are obtained with polymeric solutions of 0.4% hydroxyethylcellulose. These size pores are sufficient for sep-

6.7 Operating Characteristics of Polymer Networks 253

Dilute Semi-Dilute Entangled

0<<l)* # - 0 * (D>a)* FIGURE 6.5 Schematic representation of the entanglement process in polymer solutions, where 0 is the polymer concentration and 0* is the entanglement threshold. Reprinted with permission from Electrophoresis, 18, 2243 (1997), copyright © 1997 Wiley VCH.

arating DNA restriction fragments and polymerase chain reaction products (29), but they are too large for the separation of all but the largest proteins. For comparison, the pore size of 8%T linear polyacrylamide is in the range of 100-200 A. The broad distribution of pore sizes may contribute to the wide range of size selectivity of the polymer network (30).

It was calculated that polymer networks can provide narrow pores with short-chain polymers and larger pores with longer chain polymers, in both cases operating at a concentration near the overlap threshold (28). The polymer network systems in the aforementioned reference yield a resolution of 3 bp of DNA. With the advent of high-pressure pumping systems, unit base resolution is readily accomplished using a variety of high-viscosity networks. Table 6.2 presents a list of polymers employed in size separations.

6.7 OPERATING CHARACTERISTICS OF POLYMER NETWORKS

A. POLYMER CONCENTRATION

Optimizing the polymer concentration is a process of adjusting the molecular weight range and polymer concentration. Figure 6.6 shows the variation in migration time versus the number of base pairs for a 1-kbp DNA sizing ladder at methylcellulose concentrations of 0.2-0.6% (30). At the higher concentrations, high pressure should be used to reduce the load time. Separation between 1000 and 10,000 bp can be achieved with 0.2% MC. At 0.6% MC, resolution occurs below 100 bp. Refer to Section 5.6C for procedures to prepare low-solubility methylcellulose solutions.


Table 6.2 Materials for Replaceable Polymer Networks

Material Reference

Dextran

Galactomannan

Glucomannan

Hydroxyethylcellulose

Hydroxymethylcellulose

Hydroxypropylcellulose

Hydroxypropylmethylcellulose

Methylcellulose

Linear polyacrylamide

Polyethylene glycol

Polyethylene oxide

Polyvinyl alcohol

Polyvinylpyrrolidone

Pluronics (mixture of PEO, PPO)

Pullulan

31

32

33

28,34

27

35

29

36

37

31,32

38

32

39

40,41

42

B. INJECTION

When performing high-efficiency separations, the amount of sample injected is often the hmiting factor for resolution. The minimum amount of sample consistent with the required sensitivity should be injected. Electrokinetic injection (Section 8.3) using stacking buffers (Section 8.6) provides the best limits of detection and resolution. A comparison between electrokinetic injection and hydrodynamic injection is shown in Figure 6.7 (29). In the absence of the polymer network, all DNA or SDS-protein molecules should have the same mobility, so that electrokinetic injection is nondiscriminatory.

It is critical to desalt the sample when using electrokinetic injection (Section 8.3). The traditional means of salt removal—ethanol precipitation with recon-stitution in 50% formamide, 0.5 M EDTA—is not reproducible in terms of DNA recovery (43). Desalting is better accomplished using spin columns, at least for DNA sequencing. For many applications, simply diluting the sample is sufficient. Refer to Section 10.5 for more details.

C. FIELD STRENGTH AND TEMPERATURE

Optimization of the field strength can be performed via an Ohm's law plot (Section 2.7); however, as with polyacrylamide gels, beware of biased reptation

6.7 Operating Characteristics of Polymer Networks 255

m CD

4»» =3

03

E H-c .9 •*ri ai %m.

C ^

^

25-

20-

15-

— ' — " — I

1 2 3 4 6 Log,o Number of Base Pairs

FIGURE 6.6 Variation of the DNA fragment migration time with the log of the number of base pairs of a 1-kbp ladder using 0.2%, 0.4%, and 0.6% methylcellulose in 100 mM Tris-borate, 2 mM EDTA, pH 8, on a polyacrylamide-coated capillary. Capillary: 100 cm (80 cm to detector) x 75 |im i.d.; voltage: -30 kV; detection: UV, 260 nm. Reprinted with permission from J. Liq. Chromatogr., 15, 1063 (1992), copyright © 1992 Marcel Dekker.

effects for DNA fragments larger than 15 kbp (44). Temperature can affect DNA structure and alter mobility. For this reason, temperatures of 45°C and above are often employed to ensure denaturation. Since DNA is negatively charged at pH 8, the voltage must be set to negative polarity. MacCrehan et al. (30) found optimal resolution of the 506/517 bp fragments in 0.4% methylcellulose at 250 V/cm. The optimal efficiency was at 350 V/cm; however, the mobility differential (A/i) was superior at the lower field strength. As described by Eq. (2.17), A;U has more significance in controlling resolution.

D. USE WITH COATED CAPILLARIES

Suppression of the EOF is important when using polymer networks. While the cellulose additives do this to some extent, the EOF may still be sufficiently strong so that the solute's electrophoretic flow is overcome and larger fragments are detected first. The use of coated capillaries (Figure 6.8) reduces the EOF and provides for the normal order of migration, with smaller fragments followed by the larger ones (29, 30).

The need to control the EOF in bare silica increased the complexity of the buffers and experimental conditions. In particular, the bare silica capillary requires careful washing with hydroxide followed by equilibration with buffer. Phenylmethyl-coated

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FIGURE 6.7 Separation of a Haelll restriction digest of 0X174 DNA. Capillary: 57 cm (50 cm length to detector) x 100 |im i.d. coated with 0.1-|im-thick OV-17; buffer: 89 mM Tris-borate, 2 mM EDTA (pH 8.5), and 0.5% HPMC-4000; voltage: 10 kV; temperature: 25 °C; detection: UV, 260 nm; sample concentration: 25 |ig/mL. Key: (A) injection: electrokinetic, 5 s at 2 kV; (B) pressure injection: 60 s at 3.44 MPa. Reprinted with permission from J. Chromatogr., 559, 267 (1991), copyright 1991 Elsevier Science Publishers.

capillaries require only flushing with distilled water and methanol. When using very high viscosity polymers, the coated capillary is often unnecessary

E. PRECISION

Migration time precision (29), expressed as % relative standard deviation (%RSD), ranged from 0.09% to 0.24%. Peak height precision was 1-7%, and peak

6.8 Additional Materials for Polymer Networks 257

Migration Time in Minutes

FIGURE 6.8 Separation of a 1-kbp DNA ladder using 0.4% methylcellulose. Conditions described in Figure 6.6. Key (bp): (1) 75; (2) 134; (3) 154; (4) 201; (5) 220; (6) 298; (7) 344; (8) 396; (9) 506; (10) 517; (11) 1018; (12) 1636; (13) 2036; (14) 3054; (15) 4072; (16) 5090; (17) 6108 (18) 7126; (19) 8144; (20) 9126; (21) 10,180; (22) 11,198; (23) 12,216. Reprinted with permission from J. Liq. Chromatogr., 15, 1063 (1992), copyright © 1992 Marcel Dekker.

area precision ranged from 2% to 9%, all at DNA concentrations of 10-25 |lg/mL. At 5 |Lig/mL, precision began to deteriorate as the limit of detection was approached. Using polyacrylamide-coated capillaries (45), migration time precision was run-to-run, 0.4%; day-to-day, 0.5%; and capillary-to-capillary, 0.9%. The polyacrylamide-treated capillaries^ lasted for 50 injections before the coating deteriorated with the pH 8.0 buffer. In more recent work, a precision of 0.2 bp in 200 bases (0.1%) was found using hydroxyethylcellulose, 7 M urea, and a phenyl-methyl-coated capillary (46, 47).

6.8 ADDITIONAL MATERIALS FOR POLYMER NETWORKS

A. LINEAR POLYACRYLAMIDE

The use of linear, or uncrosslinked, polymer solutions of polyacrylamide (LPA) or dime thy Ipolyacrylamide has had profound effects, particularly in DNA sequencing. While LPA absorbs in the UV, this is not a problem for DNA separations, since LIF is usually employed.

^Bio-Rad, a manufacturer of polyacrylamide capillaries, claims to have stabilized the surface. Hundreds of injections have reportedly been obtained on a single capillary with an alkaline buffer.


Both the polymer concentration and molecular weight are important for extending the read length when sequencing. The admixture of 2% LPA (9 mDa) and 0.5% LPA (50 kDa) is used to separate DNA-sequencing reaction products of up to 1000 bases in less than 1 h (48). The viscosity of this polymer is 30,000 cps. The solution exhibits non-Newtonian properties as the viscosity drops upon the initiation of flow. The use of 2% 16-mDa LPA and 0.5% 250 kDa at 125 V/cm extends the read length to 1300 bases in 2 h (49). The lower field strength extends the read length by minimizing the biased reptation effect.

Linear polyacrylamide has also been used for antisense DNA separations where the matrix was 18%T (50), genetic analysis (51), and plasmid mapping (52).

B. DEXTRANS

Dextrans are branched polysaccharides produced by bacteria growing on a sucrose substrate. They form viscous slimy solutions when dissolved in water. Unlike agarose, which is helical in structure, dextran pores are defined by polymeric entanglement.

This material overcomes many limitations of polyacrylamide for the separation of proteins. Besides being pumpable and replaceable, the solution has a low absorbance in the low Uy permitting sensitive detection (31). Polyacrylamide-filled capillaries are usually monitored at higher wavelengths—260 nm for DNA and oligonucleotides, 280 nm for proteins. While these wavelengths are suitable for oligonucleotides, the molar absorbtivities are poor for proteins at 280 nm.

Proteins can be size separated using an electrolyte containing 0.1% SDS, 10% dextran 2 M (molecular weight 2 mDa) in 60 mM 2-amino-2-methyl-l,3-propanediol (AMPD)-cacodylic acid (CACO), pH 8.8 (Figure 6.9) (31).

Small peptides can be separated in dextran matrices as well. Using an electrolyte containing 12% dextran 2 M, 400 mM Tris-borate, 0.1% SDS, and 10% glycerol, pH 8.3, resolved myoglobin fragments ranging in molecular weight from 2515 to 16,949 on bare siUca (53). Lower concentrations of dextrans provided less-adequate resolution. Using dextran 70 K, a lower molecular weight fraction, succeeded in separating the small peptides but was inadequate for the larger fractions. The addition of glycerol increased viscosity and thus lowered the diffusion coefficients of the peptides. This is important for the smaller peptides in particular. Buffer concentrations below 200 mM Tris gave inadequate resolution.

A calibration plot of log Mj. (molecular weight) versus the migration time deviated from linearity below molecular weight 6000. This was probably due to the poor SDS-binding capacity of the small peptides. In any event, separation was achieved and was consistent with the slab gel.

Aminodextran (AD) has been used in conjunction with linear polyacrylamide (LPA) to enhance the separation of oligosaccharides (54). In this example, ion pairing between the AD (10 kDa) and negatively charged oligosaccharide

6.8 Additional Materials for Polymer Networks 259

0.012

0.0081

C o

is

< 0.004

0,000 0.00 4.00

flm« (mln) 8,00

FIGURE 6.9 Separation of standard SDS-protein complexes in a dextran polymer network. Capillary: dextran-grafted, polyacrylamide-coated 23 cm (18 cm length to detector) x 75 JLim i.d.; buffer: 60 mM AMPD-CACO, pH 8.8, 1% SDS, with 10% w/v dextran 2 M; field strength: 400 V/cm; current: 30 jiA; injection: 2 s at 300 V/cm; detection: UV, 214 nm. Key: (1) myoglobin; (2) carbonic anhydrase; (3) ovalbumin; (4) bovine serum albumin; (5) ^-galactosidase; (6) myosin. Reprinted with permission from And. Chem., 64, 2665 (1992), copyright © 1992 Am. Chem. Soc.

enhances the separation. The concentration of AD is quite low, 50 mM, so that the sieving is provided by the LPA.

C. POLYETHYLENE GLYCOL

Polyethylene glycol (PEG), a linear polymer, can also be employed as a polymer network. A 3% solution of PEG (MW 100,000) can separate SDS-proteins from 14 to 94 kDa, with detection at 214 nm (31). Protein separations are comparable with those by PEG, but the run times are somewhat longer compared with those of dextrans (Figure 6.10). Migration time RSDs of 0.4-0.5% are found when the PEG solution is replaced after each run. Ferguson plots (Section 6.1 IF) are linear and intercept they axis at the CZE mobihty values. This indicates a true size separation. More than 300 injections were performed without degradation in performance.


^ 2

OG

IAJ \] L OJ

FIGURE 6.10 Separation of low-molecular-weight standard SDS-proteins with a PEG polymer network. Capillary: dextran-grafted, polyacrylamide-coated 46 cm (40 cm length to detector) x 100 |im i.d.; buffer: 100 mM Tris-Ches, 0.1% SDS, pH 8.8, with 3% w/v PEG 100,000; field strength: 300 V/cm; injection: pressure (0.5 psig) for 20 s; detection: UV, 214 nm. Key: OG (internal standard, Orange G); (1) a-lactalbumin; (2) ovalbumin; (3) carbonic anhydrase; (4) ovalbumin; (5) bovine serum albumin; (6) phosphorylase B. Reprinted with permission from Anal. Chem., 64, 2665 (1992), copyright © 1992 Am. Chem. Soc.

D. OTHER POLYMERIC REAGENTS

It seems amazing that so many polymers can provide for size separations. Short tandem repeats have been separated using a mixture of polyethylene glycol 8 M and 350 K in TBE containing 3.5 M urea (55). Polyethylene glycols have also been used for DNA sequencing (56, 57) and protein separations (38, 58). Polyvinylpyrrolidone (39) and Pluronics (41) have been utilized for DNA

6.9 Detection 261

sequencing and oligonucleotide separations, respectively. Polyvinyl alcohol has been used for SDS-proteins (59). It is only a matter of time before designer polymers engineered for separation and detection become available.

E. COMMERCIALLY AVAILABLE GELS AND POLYMER

NETWORK REAGENTS

Both gel-filled capillaries and reagents for polymer network separations are commercially available. Table 6.3 contains a compilation of available material. Unfortunately, due to the competitive marketplace, many manufacturers choose not to reveal their specific recipes. It is likely, though, that many of these formulas are composed from reagents that have been described in this chapter.

6.9 DETECTION

The details concerning detection are covered in Chapter 9. For proteins, UV detection at the lowest possible wavelength is best. This is determined by the UV cutoff of the polymer network. For this reason, polyacrylamide is not very useful for separating SDS-proteins. When using the Bio-Rad SDS-protein matrix, a wavelength of 220 nm is best.

D N A molecules absorb strongly at 260 nm. For high sensitivity, LIF detection is always employed. Fluorophores are provided either by a fluorescent label, by a fluorescently labeled primer, by a fluorescently labeled dideoxynucleotide triphosphate, or through the use of intercalating reagents.

Intercalating reagents^ are additives to polymer network or gel systems that form complexes with specific solutes, usually DNA. These additives are employed to alter selectivity and improve detection. For example, the bands comprising 271 and 281 bp are not resolved using the buffer recipe given in Figure 6.7. If 10 |iM ethidium bromide (FtBr) is added to the buffer, the bands are completely resolved, as shown in Figure 6.11, though the run time increases by 40% (29).

Ethidium bromide is an intercalating reagent that inserts between base pairs of the DNA double helix. Since the reagent is cationic, the mobility of the DNA-EtBr complex decreases due to the reduction of the ionic charge. Incorporation of 1 |ig/mL of EtBr into a linear polyacrylamide matrix improves the separation and also the detector sensitivity (60). The UV absorbance values in the presence of EtBr are enhanced two- to threefold because of the stronger absorptivity of the DNA-EtBr complex. Since EtBr intercalates between the base

^Intercalating agents are usually toxic.

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6.9 Detection 263

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FIGURE 6.11 Effect of ethidium bromide on DNA restriction fragment resolution. Conditions as per Figure 6.7, except sample concentration: 10 }Xg/mL; injection: 2 kV for 10 s; buffer: as per Figure 5.8 with the addition of 10 jig/mL EtBr. Reprinted with permission from J. Chromatogr., 559, 267 (1991), copyright © 1991 Elsevier Science Publishers.

pairs of double-stranded DNA, there is no improvement in the separation of single-stranded oligonucleotides.

Ethidium bromide becomes planar and fluoresces strongly when intercalated in the DNA matrix. In the bulk aqueous buffer, residual fluorescence is quenched through collisions with solvent molecules. EtBr is not often used anymore in HPCE, because the absorption spectrum of EtBr does not match the emission lines of the low-cost and reliable argon-ion laser.

Separations of restriction fragments and PCR products using thiazole orange as the intercalator with 0.5% HPMC in a buffer consisting of 89 mM Tris-borate, 2 mM EDTA, pH 8.5, and 0.1-1 |Lig/mL thiazole orange resolves this problem


(61). Since the dye absorbs strongly at 488 nm, the argon-ion laser is an optimal excitation source. The limit of detection is improved by a factor of 400 compared with that of UV detection. Unlike EtBr complexes, the DNA-thiazole orange complex is sensitive to the DNA-dye concentration ratio. Good peak shapes are found when a 9:1 molar ratio of DNA:dye is employed.

Other dyes such as YO-PRO (62), 9-aminoacridine, YOYO (oxazole yellow homodimer), TOTO (thiazole yellow homodimer), and propidium also intercalate between the base pairs (63). The use of bis intercalators (homodimers) can give rise to artifact peaks from the formation (presumably) of intermolecu-lar dimers not seen when using the monomer dyes (63). SYBR Green 1 was used in conjunction with 0.3% HEC for mutation screening (64). The dye YO-PRO-1 with 0.75% HEC permitted sensitive detection of restriction fragments and PCR-amplified short tandem repeats (65). It was also shown that the accuracy of molecular weight measurements is improved when intercalators are used (66).

In a related development, hydrophobic dyes can be used to enhance detection of proteins (67). While the reported separations were by CZE, it is likely that such a scheme can be employed in polymer networks. Dyes such as 1-anilino-naphthalene-8-sulfonate (ANS) and 2-p-toluidinonaphthalene-6-sulfonate (TNS) fluoresce only when bound to a protein. Incorporation of 200 |LiM TNS in the run buffer optimized the sensitivity for conalbumin with helium-cadmium laser-induced fluorescence detection. A limit of detection of 360 nM was reported.

6.10 OPERATING HINTS USING POLYMER NETWORKS

A. FILLING THE CAPILLARY WITH POLYMER SOLUTION

Unlike a conventional BGE, the polymer networks are extremely viscous solutions. It is generally best to backfill the capillary with the polymer network to avoid coating the outside of the inlet portion of the capillary with a solution difficult to remove. If your instrument has the capability, place the polymer solution at the outlet side and pressurize the outlet vial. Operation at few hundred pounds per square inch will speed the filling of the capillary.

If you must forward-fill the capillary, then designate several rinse steps to cleanse the outer capillary wall. A rinse step should be designated even if the capillary is backfilled.

B. INJECTION

Electrokinetic injection will provide the best sensitivity and resolution, provided the salt concentration in the sample is low. If the sample contains more than 50 mM salt, hydrodynamic injection should prove superior.


C. RUN BUFFERS

The inlet and outlet reservoirs contain buffer, but there is no need for the polymer reagent to be present here. This permits the designation of one vial to contain the polymer network. That vial can be reused many times, saving valuable reagent. The same holds true when using DNA intercalators. If purchased in kit form, these reagents are expensive.

Determine experimentally how often the inlet and outlet buffers need to be changed. Refer to Section 2.1 for a discussion on buffer depletion.

D. ELECTRIC FIELD

Operate using reversed polarity. Both SDS-proteins and DNA are anionic under typical conditions and migrate toward the positive electrode. The EOF is usually nullified by the viscosity of the polymer network.

E. OPERATION AT ELEVATED PRESSURE

Occasional problems with outgassing have been observed when using viscous polymer networks. Operation of the instrument in the capillary electrochro-matography (CEC) mode permits the pressurization of the entire capillary. The use of 2-bar pressure eliminated the outgassing problem (68). The impact on migration time precision has not be adequately studied.

E CAPILLARY REGENERATION PROCEDURE

If bare silica capillaries are employed, 0.1 N sodium hydroxide and/or 0.1 N hydrochloric acid can be used as interrun washes, after which the capillary is refilled with polymer network. If a coated capillary is used, the base and usually the acid wash should be omitted. Acid washes have been used to reduce the EOF in bare silica capillaries.

6.11 APPLICATIONS AND METHODS DEVELOPMENT

The manufacturers' kits and protocols provide an appropriate starting point for most methods. The purpose of this section is to provide a basis for methods development for those wishing to develop their own separations or to adapt the kits for specific appUcations. Key apphcations will be surveyed as well. Table 6.4 presents a variety of applications and recipes that employ replaceable polymer networks.

266 C

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A. METHODS DEVELOPMENT USING POLYMER

NETWORKS—ANTISENSE D N A

The development of DNA therapeutics or gene therapy has been burdened by the rapid decomposition of the drug substances by exonucleases, enzymes that function to rid the bloodstream of foreign DNA. To overcome the biological response, research in the field is based on the development of compounds that will bind to target DNA or mRNA but are not recognized by the body's natural defenses. These are known as antisense oligonucleotides (ODNs). The most widely study ODNs are phosphothioates, which are modified by substituting a single sulfur atom for one oxygen on the phosphate group (70, 91).

During the course of synthesis of these compounds, failure sequences may occur. In the blood stream, catabolites may be formed. These fragments may still have biological activity; therefore, methods to separate the parent compound and its fragments are required. The method must use a replaceable polymer network, suppress the formation of secondary structures, and provide unit base resolution of mixtures of 15- to 20-mer ODNs.

A scheme for methods development that maps the pathway to the final method is given in Figure 6.12 (70). Among the parameters to be studied are:

1. Polymer type, molecular weight, and concentration 2. Buffers and denaturants 3. Capillary coating 4. Operational parameters, voltage, temperature, and so forth

Polymer Mesh size

Viscosity

PEG.HEC.etc.

polymer cone.

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Derivatized column

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Buffer composition!

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Buffer capacity pH-pKa

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13% PEG

Derivatized column

30% Formamide

sot lOOmM Tris-borate, pH=9.0

FIGURE 6.12 Methods development scheme employing a polymer network for the separation of antisense DNA. Reprinted with permission from Electrophoresis, 19, in press (1999), copyright 1999 © Wiley-VCH.

The selection of the polymer solution is based in part on the viscosity of the reagent blend. It must be pumpable with the instrument at hand. Figure 6.13 shows the separation in various amounts of PEG 20,000 (70). As the reagent concentration is increased, separation improves, with 13% being optimal. Above 13%, peak splitting is observed for no apparent reason. The entanglement threshold, 0*, is estimated at 1.7%, and the pore size is 20 A at 13% concentration.

A denaturing buffer system under alkaline conditions is best to suppress inter-or intramolecular hydrogen bonding of ODNs, but a pH of greater than 10 was not considered because of complications from the ionization of the T and G bases, which have pK^ values of around 10. Urea was not useful as a denaturant, but the impact of formamide, shown in Figure 6.14, is substantial (70). With the selection of pH 9 for the buffer, the potential for capillary coating degradation is significant. While good capillary stability was reported, a homemade capillary was employed, and identical results may not be found on other capillaries.

The effect of temperature is illustrated in Figure 6.15 (70). By elevating the temperature, shorter run times occur, since the viscosity of the polymer network is decreased. This in turn increases the solute mobility in the same manner as in CZE. Note the improvement in resolution at the higher temperatures. This is related to maintaining a more denaturating environment. These results

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FIGURE 6.13 Impact of PEG concentration on the separation of 15- to 20-mer antisense DNA. Capillary: polyacrylamide-coated 67 cm (60 cm length to detector) x 50 |Lim i.d.; buffer: 100 mM Tris-borate, pH 9.0, 30% formamide with the indicated amounts of PEG; voltage: -30 kV; injection: pressure (0.5 psig) for 40 s; detection: UV, 254 nm. Reprinted with permission from Electrophoresis, 19, in press (1999), copyright 1999 © Wiley-VCH.

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FIGURE 6.14 Effect on the formamide concentrat ion on resolution. Condit ions as per Figure

6.13. PEG concentration: 13%; formamide concentrations as indicated. Reprinted with permission

from Electrophoresis, 19, in press (1999), copyright 1999 © Wiley-VCH.

are all consistent with the experimental requirements for long read lengths in DNA sequencing.

B. SDS-PROTEINS

The advantages of separating SDS-proteins by capillary electrophoresis are compelling:

1. SDS solubilizes all proteins, even hydrophobic ones (13). 2. SDS-protein complexes are highly charged and very mobile (13).


3. Since the complexes are always negatively charged, they always move toward the anode (13). This holds true even if bare sihca capillaries are employed, since the SDS-protein complex is repelled from the negatively charged wall.

4. The proteins are unfolded and stretched by SDS binding (13). 5. Separation is based only on molecular weight (13). 6. Glycoforms and isoenzymes appear as a single peak, as microhetero-

geneity is not visualized (13). 7. Separation in the viscous polymer network limits diffusion, thereby opti

mizing band broadening.

1. Preparing the SDS-Protein Complex

Before performing electrophoresis, all proteins must be converted into SDS-pro-teins. This process masks individual charge differences between proteins, cleaves hydrogen bonds, cancels hydrophobic interactions, prevents aggregation, and removes secondary structure by unfolding (13).

The binding buffer usually contains 0.1% SDS and Tris, pH 9.2,5 though pH 6.8 can also be used. At this surfactant concentration, the protein concentration

5Bio-Rad SDS Sample buffer.

23t:

35T

TimeCmin)

FIGURE 6.15 Impact of temperature on resolution and speed of analysis. Conditions as per Figure 6.13. PEG concentration: 13%; formamide concentration: 30%; temperature as indicated. Reprinted with permission from Electrophoresis, 19, in press (1999), copyright 1999 © Wiley-VCH.


should be less than 1 mg/mL to ensure complete binding. The salt concentration in the sample should be kept as low as practical, especially if electrokinetic injection is employed. Hydrodynamic injection is less sensitive to salt, though band broadening may occur at concentrations above 100 mM. If the sample contains potassium, precipitation may occur through ion exchange with SDS.

Denaturation occurs by heating the protein-SDS solution in a water bath maintained at a temperature near boiling for 5-10 min. For proteins subject to fragmentation, both the time and temperature may have to be reduced. In a nonreducing environment, the sample can be left for 30 min at room temperature to minimize fragmentation. If the proteins are to be separated in the reduced forms, either 15 mM dithiothreitol (less odorous) or mercaptoethanol should be added to the buffer prior to heating to cleave the disulfide bonds.

Some proteins exhibit nonideal behavior during SDS-PAGE. Glycoproteins and lipoproteins do not bind the same amounts of SDS per gram of protein. The correlation of mobility to molecular weight falls apart under those circumstances. Aggressive denaturing conditions may be required to alleviate this problem (92). For glycoproteins, a Tris-borate-EDTA buffer increases the negative charge on the carbohydrate moiety, which does not bind SDS (13). While not studied by HPCE, it was reported that acidic and very basic nucleoproteins do not bind SDS. Alternatively, a cationic surfactant such as CTAB can be used for binding proteins at pH 3-5 (13).

2. Gels and Polymer Networks

While there have been reports on the use of rigid gels for size separation of SDS-proteins, all modern work uses replaceable polymer networks. Some of this early work established the potential of HPCE to replace the traditional slab gel. For example, the relationship between the mobiUty of each fragment and the log molecular weights was found to be linear (2). Larger proteins such as pepsin (MW 34,700) are better separated on a more porous 10%T, 3.3%C gel. Proteins always migrate faster in more porous (lower %T) gels. The same effects are observed in polymer networks.

The limitations of rigid gels are overcome by using UV-transparent and pumpable denaturing polymer networks. Proprietary formulations are being marketed by Bio-Rad and Beckman. The Beckman process has been correlated with conventional slab-gel electrophoreis for more than 50 proteins. The linear dynamic range is from a few |ig/mL to 1 mg/mL, and the molecular weight linear range is from 14 to 205 kDa. In addition, Beckman has shown migration time RSD of 0.28-0.38% and peak area RSDs of 0.87-7.0%, externally standardized. The Bio-Rad Kit provides similar specifications.

Validation of the size separation is assessed with a Ferguson plot (see Section 6.1 IF for more details). These plots are difficult to perform by HPCE with rigid gels, since at least three different %T gels in separate capillaries are required. In

the capillary format using pumpable reagents, the process is simple to implement (38). When properly denatured, all proteins should have identical charge-to-mass ratios. This is confirmed, since the Ferguson plot (data not shown) indicates all of the myoglobin proteins converging to the same point on the y axis at 0%T (2). Data comparing HPCE with the slab gel have been reported for 65 proteins with polyethylene oxide as the polymer network (93). It was concluded that both techniques yielded similar separations results and precision.

The applicability of polymer networks for performing size separations is illustrated in Figure 6.16. Data generated from the separation of a crude catalyse

i iiWll |SS

• • ft

as a 1 a 1

t4^^««t'«iiiii

[ w^:BJhm.<^imsm.mmnM 1 DeBsiixieMe Scaxi (5 iig) ^

FIGURE 6.16 Size separation of a crude denatured (boiled for 15 min in 1% SDS and 1% mercap-toethanol) catalyse preparation (1 mg/mL) by HPCE (upper trace) and slab-gel electrophoresis (lower trace). The lower trace is densimetric scan of a 10%T, 2.6%C gel. Courtesy of Applied Biosystems.


preparation are compared for runs by HPCE and a densitometer scan of a slab gel. With the x axis approximately normalized for molecular weight, a remarkable correlation between these two techniques is observed. Identification of protein dimers, trimers, and so forth, deglycosylation, and other size-modifying chemistries are the usual applications for this technique.

Many of the polymers used for size separations can bind SDS (12). When this occurs, it is possible that the polymer will compete with the protein for the surfactant or that the SDS-protein may bind to the polymer. Both polyethylene oxide and polyvinyl alcohol bind the SDS-protein in a similar fashion to their binding of the unfolded protein. In this regard, polyacrylamide and dextran are optimal, with dextran being favored because of its low UV cutoff (12).

3. Detection

Since UV detection is less sensitive than silver staining on a slab gel, there have been a few applications employing LIF detection to improve the sensitivity of SDS-protein separations. These include precapillary derivatization (94, 95) and detection of the immunoconjugate of a monoclonal antibody, v^th the naturally fluorescing drug doxorubicin (96).

4. Native Separations

There have been even fewer reported applications employing nondenaturing conditions (97). It is possible in practice to use polymer networks with amphoteric buffers such as HEPES, MES, or MOPS in the absence of SDS to observe native proteins.

C. OLIGONUCLEOTIDES

Oligonucleotides are single-stranded fragments of DNA and, as a result, do not bind to intercalators. Unless the molecules are tagged, UV detection at 260 nm is usually employed. Spectacular separations of synthetic oligonucleotides yielding millions of theoretical plates have been reported on linear polyacrylamide as well as on crosslinked gels (18, 20, 98-102).

The separation shown in Figure 6.4 of poly(dA)2o and poly(dA)4o_6o run in a 9%T linear polyacrylamide filled capillary yields unit base resolution and shows partial separation of the phosphorylated and dephosphorylated oligonucleotides. This can be important when performing drug stability studies, since dephosphorylation is one of the modes of DNA degradation. These polymer networks are prepared in situ, and today a pumpable viscous network would be


used instead. HPCE is particularly valuable for identifying failure sequences that occur during the synthesis of oligonucleotides.

Both native and denaturing gels for the separation of oligonucleotides have been studied (98). In native gels, the relative migration order is not constant for homooligomers; it depends on the base number. For base numbers less than 14, the migration order is A > C > G > T; for greater than 18 bases, the order becomes G > A > C > T. The authors suspect this discrepancy is caused by molecular bending due to self-association of guanosine. In denaturing systems, the order of migration is the same regardless of the oligonucleotide sequence.

A comparison between HPLC and HPCE for poly(dA) standards showed compelling advantages for capillary electrophoresis (21). By HPCE on a mixed-mode Neosorb-LC-N-7R column, 1-70 mer were separated with unit base resolution by a gradient run of 150 min, generating 10,000 theoretical plates. By HPCE, on a 5%T, 5%C gel at 200 V/cm, 6-255 mer were separated in 62 min, generating 2,300,000 plates (7 million plates/meter). For oligonucletides containing less than 30 bases, HPLC generally provides adequate speed and resolution. For larger oligonucletides, HPCE provides substantial separation advantages. The data in Table 6.5 provide an effective means of selecting either HPLC or HPCE as the separation tool (21).

Table 6.5 Comparative Performance of HPLC and CGE

Method

Ion-exchange HPLC Partisil SAX Nucleogen-DMA-60 MonoQ Gen-Pak FAX TSK gel DEAE-NPR

Reversed-phase LC Zorbax ODS jiBondapak C g

Mixed-mode LC Neosorb-LC-N-7R

CGE

Separated Polynucleotides

1-30 mer 1-37 mer 1-27 mer

40-60 mer 20-70 mer

2-10 mer 1-19 mer

1-75 mer

20-160 mer 19-330 mer 19-300 mer 19-340 mer

1-430 mer

Analysis Time (min)

50 110

15 39 17

25 60

95

25 70

115 70

130

Reprinted in part from J. Chromatogr., 1991; 558:273.


D. BLOTTING

Blotting involves the electrophoretic transfer of material from the gel matrix onto a membrane. Hybridization reactions using DNA probes can be used to identify the blotted material. As an alternative to traditional blotting, it is possible to perform a precapillary hybridization, with subsequent separation of the reaction products by HPCE (79).

Figures 6.17A-C show electropherograms of Joe-labeled 17-mer sequencing primer (GTAAAACGACGGCCAGT), complementary pC2 34-mer (TCGAATTCACTGGCCGTCGTTTTACAACGTCGTC), and a mixture of the two annealed at 65°C for 10 min, then slowly cooled to room temperature in 30 min (79). The third peak in Figure 6.17C was identified as a hybrid of the two reagents by laser fluorescence as well as by thermal dissociation (Figure 6.17D).

Faster and more complete hybridization is possible by incubating in dry ice, possibly due to the lack of salt in the annealing process. Salt was eliminated from the annealing process because of the deleterious impact on injection response and reproducibility. The profound impact of salt's effect is given in Table 6.6 (79). Failure to control the salt (ionic strength) concentration causes serious quantitative problems—in this case, an inverse calibration curve.

E. DNA SEQUENCING

The human genome initiative (HGI) is a project designed to sequence the entire human genome. HGI will, according to Leroy Hood, "profoundly change the study of biology and the treatment of disease" (103). HGI proposes to map and sequence the 24 different human chromosomes, which contain approximately 100,000 genes and 3 billion bases.

The project goals are as follows (104) (resolution given in parentheses):

1. Complete a detailed human genetic map (2 Mbp). 2. Complete a physical map (0.1 Mbp). 3. Acquire the genes as clones (5 kbp). 4. Determine the complete sequence (1 bp).

Capillary electrophoresis is now the predominant technique employed in step 4 of this project. Commercially available, fully automated instruments containing 96-capillary arrays, laser-induced fluorescence detection, and sophisticated base-calling computer programs are now available. The front-end sample preparation is accomplished using robotics systems.

The task at hand is to extend the read length of the sequencer to the greatest number of DNA bases possible. This will result in reduced costs, fewer sequencing reactions, higher throughput, and easier assembly of the sequenced fragments. Among the factors required to maximize the read length are the


lo©

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Joe:: |C2

. - . . w^ - ' ^^

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FIGURE 6.17 Identification of a DNA molecule by hybridization with a fluorescence-tagged oligonucleotide probe using CGE. (A) Joe-labeled 17-mer alone [5 jiig/mL in 10 mM Tris-borate-EDTA (TBE) ]; (B) pC2 alone (5iig/mL in 10 mM TBE); (C) equal amounts of Joe-labeled primer and pC2 in 10 mM TBE, driven to complete hybridization by incubation in dry ice; (D) mixture in C heated to 65°C for 5 min. Capillary: 45 cm (25 cm to detector) x 75 |im i.d. filled with a 9%T nondenaturing linear polyacrylamide gel; buffer: 25 mM Tris-borate-EDTA, pH 8.0; injection: electrokinetic, 13.5 kV for 5 s; field strength: 300 V/cm; detection: Uy 260 nm. Reprinted with permission from J. Chromatogr., 559, 295 (1991), copyright © 1991 Elsevier Science Publishers.

proper polymer blend (48), high temperature (105), appropriate field strength, and sample cleanup including desalting and template removal (106).


Table 6.6 Effect of Buffer Concentration on Electrokinetic Injection

Restriction digest, 0X174 Hae III was separated on a 9%T nondenaturing gel. The peak heights corresponding to 234, 271, and 603 base pairs (bp) fragments were compared. More DNA was injected when the sample was diluted with distilled water. A nearly 500-fold increase in peak height was observed when the sample was diluted 1000-fold. Injection was done electrokinetically at 15 kV for 5 s.

DNA Concentration (pg/mL)

1000

500

100

20

5

Tris-HCl Concentration (mM)

10

5

1

0.2

0.05

Fragment 234 bp

1

5

33

139

436

Relative Peak Height

Fragment 271 bp

1

6

27

139

442

Fragment 603 bp

1

5

24

119

364

Reprinted with permission from J. Chromatogr., 1991; 559:295.

F. DOUBLE-STRANDED DNA

This application area involves restriction digests, PCR products, genetic and mutational screening, and short tandem repeat (STR) separations. The advantage of separating double-stranded material is the ability to use intercalating dyes to enhance detection.

1. Restriction Digests

Restriction digests are mixtures of DNA fragments produced by the reaction of DNA and a restriction enzyme, an enzyme that cuts at specific base sequences. These enzymes are used for the creation of genetic maps prior to sequencing.

An HPMC polymer network system can be used to monitor a PCR-amplified human immunodefieciency virus (HIV) infected cell line (Ul.l) (29). The cell line contains one copy of HIV-1 provirus and two copies of HLA-DQ-a, which is normally present in all healthy cells. Using specific primers, a 115-bp region of HIV-1 and a 242-bp region of HLA-DQ-a were coamplified by 35 cycles of PCR. Separations with and without ultracentrifugation are shown in Figure 6.18. The ultracentrifuge simultaneously desalts and concentrates the DNA in the sample.

Since the concentration of the polymeric additive is easily varied, generation of a Ferguson plot (log JJ. vs. %T) is simple. Plotting the HPMC-4000 concentration versus log mobility for a 0X174 Hae III restriction digest (Figure 6.19) shows imperfect convergence at 0% polymer (29).


No CrafarltSfHi

ffl?-I

Tttm (mln)

FIGURE 6.18 Separation of a PCR-amplified cell line containing HlV-1 provirus. Voltage: 20 kV; injection: electrokinetic, 10 kV for 10 s. Other conditions as per Figure 6.7. (A) no ultracentrifu-gation; (B) Ix ultracentrifugation; (C) 2x ultracentrifugation; (D) 3x ultracentrifugation; (E) 0X174 DNA standard. Reprinted with permission from J. Chromatogr., 359, 267 (1991), copyright © 1991 Elsevier Science Publishers.

There are four types of behavior expected in the Ferguson plot (13):

1. The hues are parallel. The molecules have the same size but different mobilities (e.g., isoenzymes).

2. The slopes are different, but the lines do not cross. The molecule giving the upper curve is smaller and has a higher net charge.

3. The lines cross at high polymer concentration. The larger molecule has the higher charge density.

4. Several lines cross at low polymer concentration, or they converge when extrapolated to 0% polymer. These molecules are all similar.


-3.4

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% (w/w) HPMC-4000

FIGURE 6.19 Ferguson plot (log ju vs. % w/w HPMC) for a buffer containing HPMC-4000 as the polymer network. Mobihties of selected 0X174 Hue 111 digest fragments were used to generate the plot. Key:0 = 118bp;A = 194bp;n = 310bp;+ = 603 bp; A = 872bp;# = 1353 bp. Reprinted with permission from J. Chromatogr., 559, 267 (1991), copyright © 1991 Elsevier Science Publishers.

For the Ferguson plot illustration, case 4 provides the best fit. Imperfect convergence does not necessarily mean errors will occur when calculating molecular weight. If the data for a molecule are nonlinear, this is indicative of problems such as experimental errors or adhesion of the molecule to the gel.

2. Short Tandem Repeats (STRs)

DNA is now employed for human identification for forensic and military purposes. In the early 1990s, a technique known as restriction fragment length polymorphism (RFLP) was the method of choice. The technique works based on the presence or absence of restriction sites. Problems with the technique


include the need for large amounts of intact DNA (20-100 ng) and the need for radioactive isotopes (107), although chemiluminescence is now used. The technique is time-consuming and labor-intensive. The large number of alleles that differ by only a few sequences can be difficult to separate (107).

The RFLP procedure is being replaced by a PCR method known as short tandem repeats (STRs) (46, 47, 90,108-112). Otherwise known as microsatellites, short tandem repeats are repetitive sequences where 2-7 nucleotides of DNA are repeated over and over again. Unlike DNA found in a gene, short tandem repeats are especially prone to DNA replication errors known as slip-strand mis-pairing (113). As a result, the lengths of these DNA satellites vary from one person to the next, and thus, they provide the potential for DNA fingerprinting.

The use of LIF detection provides for the high sensitivity of the method. A separation of the HUMTHOl allelic ladder in a 1% HEC network using TBE buffer and 500 ng/mL YO-PRO-1 as fluorescent intercalator in shown in Figure 6.20 (90). The size of each fragment is calculated based on the migration times of the 150-bp and 300-bp internal standards.

The need to extract more information from a sample and to preserve DNA has led to an approach known as multiplex PCR. The procedure involves adding more than one set of PCR primers in order to amplify multiple locations throughout the genome. The probability of finding identical alleles in individuals decreases as the number of loci examined are increased. Since fluorescent labeling with different dyes is used, detection of STRs with the same size range is readily accomplished. Multiple dyes also permit the simultaneous separation of a standard along with the unknown (89). This is the approach employed using the ABI Prism 310 Genetic Analyzer (110). The instrument has four-dye capability, one for the standard and three for the samples. Using that instrument, fragments of less than 350 bp can be separated in 30 min.

It is expected that capillary arrays will play a huge role in STR analysis, since high throughput is required due to the expected sample load. Microfabricated devices that integrate the PCR step with separation and detection may play a role here as well (114).

Capillary electrophoresis is an ideal technology for forensic DNA analysis, since the process is completely automated. There is no need to manually pour the gel or pipet the sample. Because of issues concerning validation, virtually all forensic laboratories will opt for a commercial instrument and protocol.

3. Genetic Analysis

Allele-specific amplification can be employed to detect a single base-pair mutation through the use of a specially designed primer complementary to the mutated DNA (64, 78). PCR amphfication only takes place if the mutation is present. HPCE of the now double-stranded material takes place in a 4% LPA matrix on an 8-cm DB-1 capillary with a buffer containing Tris-TAPS, 2 mM

<5.00 ^

40.00

35.00 -J

30.00 -J

25.00

115.00

liO.OO

105.00 ~J

100.00

8.00

150 bp

HUMTHOl

Alleles

300 bp

Ca»cylatg<i gizg

183.52 187.66 191.72 195.70 199.77 202.89 207.81

1.00 o.bo Minutes

FIGURE 6.20 Separation of short tandem repeats in a HUMTHOl allehc ladder. Capillary: DB-17 coated 27 cm (20 cm effective length) x 50 |Lim i.d.; buffer: 1% HEC, 100 mM Tris-borate, 2 mM EDTA, pH 8.2, with 500 ng/mL YO-PRO-1; temperature: 25°C; injection: electrokinetic: 1 kV for 5 s; voltage: 5 kV; detection: LIF; excitation at 488 nm, emission at 520 nm. Reprinted with permission from J. Chromatogr., B: Biomed. Appl 658, 271 (1994), copyright © 1994 Elsevier Science Publishers.

EDTA, pH 8.3, carrying 0.5 |Ig/mL EtBr,using LIF with a doubled Nd-YV04 laser (78).

The most widely used genetic screening technique, PCR-RFLP, detects a mutation by the availability of a specific restriction endonuclease cleavage site at the mutation locus (115). The products can be separated with a polymer network containing an intercalator. The products from other techniques such as ARMS (amplification refractory mutation system), SSCP (single-strand conformational polymorphism) (116, 117), HPA (heteroduplex polymorphism) (118), and CDCE (constant denaturant capillary electrophoresis) (119) are all amenable to polymer networks.


In CDCE, separation occurs takes place in a heated portion of the capillary, where faster moving, unmelted DNA fragments are in equilibrium with slower moving partially melted forms (120). Depending on the temperature range, the melting equilibrium and the average mobility of each mutant gene is different, and single base-pair point mutations can be resolved. The mobility is highest when DNA is in its double-stranded form at lower temperatures (e.g., 45°C), the exact temperature depending on the DNA sequence. Mutants are not resolved under these conditions. At high temperatures (e.g., 51°C), the now single-stranded mutants are not well separated. At intermediate temperatures, separations with unit base resolution can be observed. A typical BGE is 5% LPA in 1 X TBE using an LPA-coated capillary.

G. PLASMIDS

Plasmids are circular bacterial DNA fragments that can exist in twisted super-coiled forms. They have the properties of a virus, but without the outer protein membrane. Several reports describing plasmid separations by HPCE have appeared in the literature (121, 122). Polymer networks are best used for plasmid separations. Separations using 0.1% HEC, 0.1% HPMC, or 0.15% PEO all in 1 X TBE provide good separations using coated capillaries (123).

H. RNA

RNA separations are performed to determine RNA mass and conformation in the determination of gene expression, in the identification of microorganisms including retroviruses and bacteria (124), and from paraffin-embedded postmortem tissue specimens amplified by PCR (125). To minimize RNA degradation, material should be kept at -70°C until analysis (125). In some cases, RNA is detected via PCR that provides cDNA, which is further amplified, separated, and detected (125). For high-sensitivity detection of cDNA, the use of interca-lators and laser-induced fluorescence (LIE) is indicated. A Beckman dsDNA 1000 Kit, which contains a 10% LPA capillary in TBE and 0.4 |Lig/mL EnhanCE intercalator (125) or 1% HEC in TBE with 1 |iM YO-PRO on a DB-17 capillary (125), can be used in conjunction with the argon-ion laser.

Since RNA is single-stranded, the use of intercalators is not possible, and UV detection must be employed. Many of the same polymer networks used for DNA are appUcable to RNA. The migration behavior of RNA is similar to single-stranded DNA, though RNA becomes slightly less mobile at lengths over 1000 bases. A typical electrolyte system is 0.3-0.7% HPCE (4000 cp, 2% solution) in TBE buffer containing 8 M urea. Separations are best performed at high temperatures (50°C


or higher) to shorten the run time and maintain the denaturing environment. Resolution is lost when the field strength is over 300 V/cm. Sample preparation involves, at a minimum, dissolving in 80% formamide and heating to 95°C for 3 min to ensure denaturation (124).

6.12 REDUCING THE PROBLEM OF BIASED REPTATION

Biased reptation limits the DNA fragment size that can be separated using a continuous electric field. In the absence of the field, DNA and large oligosaccharides as well are tightly coiled. When the voltage is applied, the molecule lengthens as it aligns with the field. Beyond 20 kbp, fragments are no longer resolved by constant-field electrophoresis.

Two techniques can be used to extend the range of separation for molecules following the biased reptation mechanism: voltage gradients (126) and pulsed-field electrophoresis (127-129). Voltage gradients can be performed on most HPCE instruments. No pulsed-field instruments are commercially available.

A. VOLTAGE GRADIENTS

At a constant field of 200 V/cm, all 0X174 restriction fragments are separated, but the run time is 27 min (Figure 6.21A). Increasing the field strength speeds the separation at the expense of resolution for the larger fragments. By running a voltage ramp starting at 400 V/cm and decreasing to 100 V/cm over 10 min, all fragments are resolved in less than 10 min (Figure 6.2IB) (126). The small fragments are not strongly aligned with the field and thus can be exposed to the high field strength. By the time the larger fragments are ready to elute, the field strength is sufficiently low that separation occurs.

This method is not employed in DNA sequencing instruments, since the timing of peak elution would change with time. While it would probably extend the read length, the additional complexity added in the algorithms may decrease the reliability of the base calling. The technique has not been widely utilized for other applications, since the reproducibility may be worse than with constant voltage.

B. PULSED-FlELD ELECTROPHORESIS

Pulsed-field electrophoresis has been used for some time for the separation of large DNA fragments in the slab gel (130). In its simplest format, the electric field driving the separation is pulsed. During the off cycle, the DNA molecules

6.12 Reducing the Problem of Biased Reptation 285

mm S

FIGURE 6.21 Separation of 0X174 DNA restriction fragments by CGE using (A) a constant applied electric field of 200 V/cm and (B) a linear field strength gradient ramping from 400 to 100 V/cm in 20 min. Capillary: polyacrylamide gel-filled DB-225 coated 40 cm (length to detector, 27 cm) X 100 |xm i.d.; buffer: 100 mM Tris-borate, 2 mM EDTA, pH 8.35. Key (bp): (1) 72; (2) 118; (3) 194; (4) 234; (5) 271; (6) 281; (7) 310; (8) 603; (9) 872; (10) 1078; (11) 1353. Reprinted with permission from Anal Chem., 64, 2348 (1992), copyright © Am. Chem. Soc.

relax. Upon return of the field, molecules begin migration in a partially deformed state, since alignment with the field is a kinetic process. This mechanism provides additional selectivity for the separation of large fragments.


Alternatively, multiple-electrode systems that provide two electric fields can be employed. For example, electrodes can be positioned at angles of 90 or 120°. The applied voltage can be alternated between the electrode sets at frequencies designed to enhance selectivity for various size ranges. Upon application of the pulse, DNA molecules reorient in the gel. Longer fragments reorient slowly, and their migration through the gel is hindered. Isolation and mapping of large segments of genomic DNA is an important application of this technique. Separations of several million base pairs are possible using pulsed-field techniques.

Adaptation of this technique to HPCE is in its infancy. An instrument was developed to deliver the pulsed field in either the unidirectional (single-polarity) or field-inversion (polarity-reversed pulse) formats, which position the electrodes at a relative angle of 180° (9, 37). Other angular placements used in slab-gel electrophoresis are not compatible in capillaries. Waveform distortion due to the high resistance of the gel-buffer system limited the pulse frequency to less than 100 Hz. While this problem may be solved by increasing ionic strength and using active cooling to remove heat, commercialization of this technique is probably years away.

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82. Hammond, R. W, Oana, H., Schwinefus, J. J., Bonadio, J., Levy, R. J., Morris, M. D. Capillary Electrophoresis of Supercoiled and Linear DNA in Dilute Hyrdoxyethyl Cellulose Solution. Anal. Chem., 1997; 69:1192.

83. Rill, R. L., Liu, Y., Van Winkle, D. H., Locke, B. R. Pluoronic Copolymer Liquid Crystals: Unique Replaceable Media for Capillary Gel Electrophoresis. J. Chromatogr, A, 1998; 817:287.

84. Tsuji, K. High-Performance Capillary Electrophoresis of Proteins. Sodium Dodecyl Sul-phate-Polyacrylamide Gel-Filled Capillary Column for the Determination of Recombinant Biotechnology-Derived Proteins. J. Chromatogr, 1991; 550:823.

85. Kleemiss, M. H., Gilges, M., Schomburg, G. Capillary Electrophoresis of DNA Restriction Fragments with Solutions of Entangled Polymers. Electrophoresis, 1993; 14:515.

86. McGregor, D. A., Yeung, E. S. Detection of DNA Fragments Separated by Capillary Electrophoresis Based on Their Native Fluorescence inside a Sheath Flow. J. Chromatogr, A, 1994; 680:491.

87. Katsivela, E., Hofle, M. G. Low-Molecular-Mass RNA Fingerprinting of Bacteria by Capillary Electrophoresis Using Entangled Polymer Solutions. J. Chromatogr, A, 1995; 717:91.

88. Stanta, G., Bonin, S. RNA Quantitative Analysis from Fixed and Parraffin-Embedded Tissues: Membrane Hybridization and Capillary Electrophoresis. BioTechniques, 1998; 24:271.

89. Wang, Y, Ju, J., Carpenter, B. A., Atherton, J. M., Sensabaugh, G. E, Mathies, R. A. Rapid Sizing of Short Tandem Repeat Alleles Using Capillary Array Electrophoresis and Energy-Transfer Fluorescent Primers. Anal. Chem., 1995; 67:1197.

90. Butler, J. M., McCord, B. R., Jung, J. M., Wilson, M. R., Budowle, B., Allen, R. O. Quantitation of Polymerase Chain Reaction Products by Capillary Electrophoresis Using Laser Fluorescence. J. Chromatogr B: Biomed. Appl, 1994; 658:271.

91. Maier, M., Fritz, H., Gerster, M., Schewitz, J., Bayer, E. Quantitation of Phosphorothioate Oligonucleotides in Human Blood Plasma Using a Nanoparticle-Based Method for Solid-Phase Extraction. Anal. Chem., 1998; 70:2197.

92. Dunn, M. J. Electrophoresis in Polyacrylamide Gels, in New Directions in Electrophoretic Methods, Eds. J. W Jorgenson and M. Phillips. 1987, American Chemical Society. 20.

93. Guttman, A., Nolan, J. Comparison of the Separation of Proteins by Sodium Dodecyl Sul-fate-Slab Gel Electrophoresis and Capillary Sodium Dodecyl Sulfate-Gel Electrophoresis. Anal. Biochem., 1994; 221:285.

References 291

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95. Gump, E. L., Monnig, C. A. Pre-column Derivatization of Proteins to Enhance Detection Sensitivity for Sodium Dodecyl Sulfate Non-gel Sieving Capillary Electrophoresis. J. Chromatogr., A, 1995; 715:167.

96. Liu, J., Abid, S., Lee, M. S. Analysis of Monoclonal Antibody Chimeric BR96-Doxorubicin Immunoconjugate by Sodium Dodecyl Sulfate-Capillary Gel Electrophoresis with Ultraviolet and Laser-Induced Fluorescence Detection. Anal. Biochem., 1995; 229:221.

97. Zhu, M., Levi, V, Wehr, T. Size-Based Separations of Native Proteins and SDS-Protein Complexes Using Nongel Sieving Capillary Electrophoresis. Amer. Biotech. Lah., 1993; 11:26.

98. Guttman, A., Nelson, R. J., Cooke, N. Prediction of Migration Behavior of Oligonucleotides in Capillary Gel Electrophoresis. J. Chromatogr., 1992; 593:297.

99. Cohen, A. S., Najarian, D. R., Paulus, A., Guttman, A., Smith, J. A., Karger, B. L. Rapid Separation and Purification of Oligonucleotides by High-Performance Capillary Gel Electrophoresis. Proc. Natl. Acad. Set, 1988; 85:9660.

100. Guttman, A., Cohen, A. S., Heiger, D. N., Karger, B. L. Analytical and Micropreparative Ultrahigh Resolution of Oligonucleotides by Polacrylamide Gel High-Performance Capillary Electrophoresis. Anal. Chem., 1990; 62:137.

101. Dubrow, R. S. Analysis of Synthetic Oligonucleotide Purity by Capillary Gel Electrophoresis. American Lah, 1991; March:64.

102. Yin, H.-E, Kleemis, M. H., Lux, J. A., Schomburg, G. Diffusion Coefficients of Oligonucleotides in Capillary Gel Electrophoresis. J. Microcolumn Sep., 1991; 3:331.

103. Newman, A. R. The Human Genome Initiative: First Steps. Anal. Chem., 1990; 62:1260. 104. Mapping and Sequencing the Human Genome.

http:www.ornl.gov/TechResourcesMummij3enome/piMicat/primer/pnrn2.html, 1999; 105. Klepernik, K., Foret, E, Berka, J., Goetzinger, W, Miller, A. W, Karger, B. L. The Use of Ele

vated Column Temperature to Extend DNA Sequencing Read Lengths in Capillary Electrophoresis with Replaceable Polymer Matrices. Electrophoresis, 1996; 17:1860.

106. Salas-Solano, O., Ruiz-Martinez, M. C, Carrilho, E., Kolter, L., Karger, B. L. A Sample Purification Method for Rugged and High-Performance DNA Sequencing by Capillary Electrophoresis Using Replaceable Polymer Solutions. B. Quantitative Determination of the Role of Sample Matrix Components on Sequencing Analysis. Anal. Chem., 1998; 70:1528.

107. Williams, P E., Marino, M. A., Del Rio, S. A., Turni, L. A., Devaney J. M. Analysis of DNA Restriction Fragments and Polymerase Chain Reaction Products by Capillary Electrophoresis. J. Chromatogr, A, 1994; 680:525.

108. Deforce, D. L. D., Millecamps, R. E. M., Van Hoofstat, D., Van den Eeckhout, E. G. Comparison of Slab Gel Electrophoresis and Capillary Electrophoresis for the Detection of the Fluo-rescently Labeled Polymerase Chain Reaction Products of Short Tandem Repeats. J. Chromatogr, A, 1998; 806:149.

109. Dimo-Simonin, N., Grange, E, Kratzer, A., Brandt-Casadevall, C , Mangrin, P Forensic Validation of the Short Tandem Repeat HUMACTBP2 Using Capillary Electrophoresis. Electrophoresis, 1998; 19:256.

110. Lazaruk, K., Walsh, P S., Oaks, E, Gilbert, D., Rosenblum, B. B., Menchen, S., Scheibler, D., Wenz, H. M., Holt, C, Wallin, J. Genotyping of Forensic Short Tandem Repeat (STR) Systems Based on Sizing Precision in a Capillary Electrophoresis Instrument. Electrophoresis, 1998; 19:86.

H I . Mansfield, E. S., Vainer, M., Enad, S., Barker, D. L., Harris, D., Rappaport, E., Fortina, P Sensitivity, Reproducibility, and Accuracy in Short Tandem Repeat Genotyping Using Capillary Array Electrophoresis. Genome Res., 1996; 6:893.

112. Vainer, M., Enad, S., Dolnik, V, Xu, D., Bashkin, J., Marsh, M., Tu, O., Harris, D. W, Barker, D. L. Short Tandem Repeat Typing by Capillary Array Electrophoresis: Comparison of Sizing Accuracy and Precision Using Different Buffer Systems. Genomics, 1997; 41:1.

113. Moxon, E. R., Wills, C. DNA MicrosateUites: Agents of Evolution. Scientific Amer, 1999; 280:94.


114. Woolley, A. T., Hadley D., Landre, P., deMello, A. J., Mathies, R. A. Functional Integration of PCR Amplification and Capillary Electrophoresis in a Microfabricated DNA Analysis Device. Anal. Chem., 1996; 68:4081.

115. Mitchelson, K. R., Cheng, J. Point Mutation Screening by High-Performance Capillary Electrophoresis. J. Capillary Electrophor., 1995; 2:137.

116. Kuypers, A. W. H. M., Willems, P M. W, van der Schans, M. J., Linssen, P C M . , Wessels, H. M. C, de Bruijn, C. H. M. M., Everaerts, E M., Mensink, E.J. B. M. Detection of Point Mutations in DNA Using Capillary Electrophoresis in a Polymer Network. J. Chromatogr., B: Bio-med.Appl, 1993; 621:149.

117. Baba, Y. Analysis of Disease-Causing Genes and DNA-Based Drugs by Capillary Electrophoresis. Towards DNA Diagnosis and Gene Therapy for Human Diseases. J. Chromatogr., B: Biomed. Appl, 1996; 687:271.

118. Cheng, J., Kasuga, T., Mitchelson, K. R., Lightly E. R., Watson, N. D., Martin, W. J., Atkinson, D. Polymerase Chain Reaction Heteroduplex Polymorphism Analysis by Entangled Solution Capillary Electrophoresis. J. Chromatogr, A, 1994; 677:169.

119. Khrapko, K., Hanekamp, J. S., Thilly W. G., Belenkii, A., Foret, E, Karger, B. L. Constant Denaturant Capillary Electrophoresis (CDCE): A High Resolution Approach to Mutational Analysis. Nucleic Acids Res., 1994; 22:364.

120. Khrapko, K., CoUer, H., Thilly, W. Efficiency of Separation of DNA Mutations by Constant Denaturant Capillary Electrophoresis Is Controlled by the Kinetics of DNA Melting Equilibrium. Electrophoresis, 1996; 17:1867.

121. Nackerdien, Z., Morris, S., Choquette, S., Ramos, B., Atha, D. Analysis of Laser-Induced Plas-mid DNA Photolysis by Capillary Electrophoresis. J. Chromatogr, B: Biomed. Appl, 1996; 683:91.

122. Courtney, B. C , Williams, K. C, Bing, Q. A., Schlager, J. J. Capillary Gel Electrophoresis as a Method to Determine Ligation Efficiency Anal. Biochem., 1995; 228:281.

123. Mao, D. T., Yu, L., Lautamo, M. A. High Resolution CE Separation of Supercoiled Plasmid DNAs and Their Conjormers in Dilute Polymer Solutions Containing No Intercalator, presented at the Frederick Conference on Capillary Electrophoresis. 1997.

124. SkeidsvoU, J., Ueland, P M. Analysis of RNA by Capillary Electrophoresis. Electrophoresis, 1996; 17:1512.

125. Borson, N. D., Strausbauch, M. A., Wettstein, R J., Oda, R. P, Johnston, S. L., Landers, J. P Direct Quantitation of RNA Transcripts by Competitive Single-Tube RT-PCR and Capillary Electrophoresis. BioTechniques, 1998; 25:130.

126. Guttman, A., Wanders, B., Cooke, N. Enhanced Separation of DNA Restriction Fragments by Capillary Gel Electrophoresis Using Field Strength Gradients. Anal. Chem., 1992; 64:2348.

127. Kim, Y, Yeung, E. S. DNA Sequencing with Pulsed-Field Capillary Electrophoresis in Poly(eth-ylene oxide) Matrix. Electrophoresis, 1997; 18:2901.

128. Kim, Y., Morris, M. D. Ultrafast High Resolution Separation of Large DNA Fragments by Pulsed-Field Capillary Electrophoresis. Electrophoresis, 1996; 152.

129. Ueda, M., Oana, H., Baba, Y, Doi, M., Yoshikawa, K. Electrophoresis of Long DNA Molecules in Linear Polyacrylamide Solutions. Biophys. Chem., 1998; 71:111.

130. Anand, R., Southern, E. M. Pulsed Field Gel Electrophoresis, in Gel Electrophoresis of Nucleic Acids: A Practical Approach, Eds. D. Rickwood and B. D. Hames. 1990, IRL Press. 101.

CHAPTER 7 Capillary Electrochromatography

7.1 Introduction 7.2 Modes of CEC 7.3 Electroosmotic Flow in CEC 7.4 Efficiency of CEC 7.5 Operating Characteristics of Packed CEC 7.6 Applications 7.7 CEC Microfluidic Devices

References

7.1 INTRODUCTION

At HPCE'95, the Seventh International Symposium on Capillary Electrophoresis, the Hewlett-Packard Corporation sponsored a workshop on capillary electrochromatography (CEC). A 15' X 25' room was reserved for the workshop. When 400 conferees appeared, two things became obvious:

1. A larger room was needed. 2. Separation scientists were intrigued or at least curious by the potential

of CEC. To quote Professor Csaba Horvath, "We are back to doing chromatography again."

Technological advances over the last century have governed the means of driving the mobile phase in chromatographic separations. Classical column chromatography relied on gravity, paper and thin-layer chromatography employed capillary action, HPLC used hydraulic pressure, and finally, MECC and CEC depend on the EOF to drive the mobile phase (1).

The first report describing CEC appeared in 1974, when Pretorius et al. (2) demonstrated the possibility of using the EOF to drive methanol.water

293

294 Chapter 7 Capillary Electrochromatography

through a 1-mm glass tube packed with 75-125 JLiin octane-coated Partisil. In this pre-Jorgenson era paper, the lack of injection and detection mechanisms prevented actual separations from being performed, but a concept was born.

In 1981, Jorgenson and Lukacs (3) produced an electrically driven separation of 9-methylanthracene from perylene in a 170-|Llm-i.d. Pyrex tube packed with 10-|im Ci8 particles. The separation efficiencies were modest—for example, 31,000 theoretical plates for perylene—and the authors said that "the performance of these columns appears to offer a modest improvement over conventional [pressure-driven] flow, but may not justify the increased difficulty in working with electroosmotic flow."

Despite the early difficulties, the prospects for employing an electroosmotic pumping system for liquid chromatography are tantalizing. The characteristics of EOF have three potential advantages over pressurized flow:

1. The facile generation of very low flow rates. The need for such low flows is established in Table 7.1, both for packed and for open tubular capillaries. Generation of EOF-pumped low flow rates is simple and can be controlled by selection of the packing material, the mobile phase, and the applied electric field.

2. The plug flow characteristics of the electrodriven system should prove advantageous compared with the laminar profile of the pressure-driven system (Section 2.3).

3. The EOF-driven system operates at atmospheric pressure and is pulse-free, an advantage when interfaced to the mass spectrometer.

The distinguishing feature separating electrophoresis from chromatography is retention. In HPLC, retention is the driving force for separation. In HPCE,

Table 7.1 Column Diameters and Nominal Flow Rates for LC and CEC

Category Column i.d. Nominal Flow Rate (pL/min)

Analytical LC

Small bore LC

Micro bore LC

Packed capillary LC

Packed capillary CEC

Open tubular LC

Open tubular CEC

Open tubular CEC

Open tubular CEC

4.6 mm

2.0 mm

1.0 mm

350 pm

170 pm

50 pm

25 pm

10 pm

5 pm

1000

189

47

5.8

1.4

0.12

0.029

0.0047

0.0012

7.2 Modes of CEC 2 9 5

retention must be avoided at all costs, or the theoretical maximum efficiency as predicted by Eq. (2.15) will never be obtained.

Retention in CEC is based on solute partitioning between the mobile and stationary phases. Charged solutes can also migrate via electrophoresis as well providing a multimechanistic retention scheme. In Chapter 4, the use of secondary equilibrium with micelles and/or cyclodextrins (CDs) for separation of neutral or charged molecules was described. The advantages CEC over MECC are:

1. Much of the vast HPLC literature may be applicable to CEC. 2. Interface to the mass spectrometer is more robust due to the absence of

surfactant. 3. Sensitivity in laser fluorescence is improved due to the lower reagent

background. 4. The ability to easily separate water-insoluble compounds. 5. The injection solvent is compatible with the mobile phase when sepa

rating hydrophobic solutes. 6. High loading capacity compared with that of CZE.

On the other hand, MECC has some significant advantages over CEC. These include:

1. The ability to use inexpensive and rugged bare silica capillaries. 2. Rapid equilibration of the capillary. 3. Powerful tools for the adjustment of selectivity.

7.2 MODES OF CEC

The are three major classes of CEC: open tubular, packed capillary, and CEC with replaceable media.

A. OPEN TUBULAR CEC

The use of a capillary coated with a stationary phase was reported in 1982 by Tsuda et al. (4). The immediate advantage of the open tubular format is the relative ease of manufacture of the capillaries. Well-understood surface chemistries can be applied, and frits are unnecessary. Difficulties in packing small-diameter capillaries are avoided as well. Bubble formation, a problem with packed capillaries, does not occur.

Tubing diameters of 5- to 50-|lm i.d. are used. Since solute diffusion to the capillary coating is required for retention, retention and efficiency both increase as the tubing diameter is decreased. On the other hand, detection


and loading capacity become problematic as the capillary diameter becomes narrow. In 50-|Lim capillaries, relatively small k' values are found (5), since the solute spends much of its time in the mobile phase.

It appears that open tubular CEC works best for charged solutes, since they can electrophorese as well. The purpose of the chromatographic interaction is utilized to affect the selectivity of the separation. Separations for aromatic hydrocarbons were reported using 10-|im capillaries, but the V values were less than 0.2 (6). Better results were found for chiral separations using immobilized cyclodextrins (7, 8), but superior separations are often found when simply adding CDs to the BGE. Separation of the enantiomers of a binaphthyl derivative is shown in Figure 7.1 (8).

Capillaries derivatized with Cjg, diol and cholesterol functionality are available from Silicon Valley Separation Media, c/o Dr. J. Pesek, San Jose State University (San Jose, CA).

'^-^ J A - J U U-

8 12 16 20 24

FIGURE 7.1 Separation of the enantiomers of l,l'-binaphthyl-2,2'-diyl-hydrogen phosphate at different apphed vokages. Capillary: Chiralsil-Dex, 80 cm effective length x 50 |Lim i.d.; buffer: borate-phosphate, pH 7; detection: UV, 220 nm; temperature: 20°C. Reprinted with permission from J. High Res. Chromatogr., 15, 129 (1992) copyright © 1992 Dr. Alfred Heuthig Publishers.

7.2 Modes of CEC

B. PACKED CEC

297

The majority of reported work has employed packed CEC. The advantages of this mode have been described Section 7.1. An illustration of a packed capillary is shown in Figure 7.2.

The total migration time t^ is (9)

(7.1) V 1 . V

packed open

where L = the length of the capillary to the detector, L^^^cked = the length of the chromatographic packing, L pen = the unpacked portion of the capillary leading to the detector, Vp cked = migration velocity through the packing, and open = migration velocity in the open segment of the capillary.

For neutral solutes, v^^^^ is defined by the EOF On the other hand, charged solutes may show differential migration in the open segment. If this occurs, the open segment may be lengthened to take advantage of this effect. Thus, the combination of chromatography and electrophoresis may improve selectivity over what is found in either mode alone (9).

Among the disadvantages of CEC are the newness of the technique, problems with the production of capillaries, frit manufacture, bubble formation in the mobile phase, the lack of gradient elution equipment on some instruments, and the tendency of amines to stick to the packing material (10).

Some of these issues are being addressed. New instruments from Unimicro Technologies (San Francisco, CA) and Micro-Tech Scientific (Sunnyvale, CA) designed for CEC were introduced at HPCE'99. The ultimate instrument would be a gradient system capable of CZE, CEC, and |i-HPLC.

The use of 2 |iL/mL of hexylamine as a mobile-phase additive reduces peak tailing from the interaction of basic compounds with silanols. Meanwhile, this additive only modestly decreases the EOF (11).

Detection Window Frit Frit

FIGURE 7.2 Diagram of a packed CEC capillary


The problem of frits can be addressed by eliminating them. The development of "monolithic" technology permits the immobilization of chromatographic media within the capillary in a fritless environment (12-14).

Since the CEC packing has such a large surface area, bubbles tend to form on the packing surface. This problem is minimized by running with both the inlet and outlet pressurized to 10 bar, the so-called "CEC mode."

C. CEC WITH REPLACEABLE MEDIA

A pumpable entangled polymer solution has been used for separation of neutral solutes using conventional HPCE instrumentation (15). The polymer was composed of 40% ethyl acrylate, 50% methacrylic acid, and 10% lauryl methacrylate. Typically, the capillary would be refilled daily, but with the advent of high-pressure instrumentation, the capillary could be refilled prior to each run if necessary. A highspeed separation of polycycHc aromatic hydrocarbons in shown in Figure 7.3 (15).

In a related development, separations of small molecules have been demonstrated in 10%T linear polyacrylamide gels (16) as well as in crosslinked gels (17). While these gels were not considered replaceable at the time, the 10%T gel can be replaced at high pressure.

While few papers have been published in this field and much work remains, it is easy to envision that this mode of CEC will become important. Consider-

FIGURE 7.3 High-speed separation of aromatic hydrocarbons using a replaceable stationary phase. Capillary: 25 cm x 50 |im (total length 33 cm) bare silica filled with 4% polymer; mobile phase: 40% acetonitrile, 10 mM borate, pH 9.2; voltage: 30 kV; injection: 1 kV for 30 s; temperature: 25°C; detection: UV, 200 nm. Key: (1) benzene; (2) unknown; (3) naphthalene; (4) fluorene; (5) anthracene; (6) pyrene; (7) chrysene; (8) benzo(e)pyrene; (9) benzo(ghi)perylene. Redrawn with permission from Anal. Chan., 70, 4985 (1998) copyright © 1998 Am. Chem. Soc.

7.3 Electroosmotic Flow in CEC 299

ing what has happened in the area of size separations, replaceable media have effectively displaced the use of rigid gels.

7.3 ELECTROOSMOTIC FLOW IN CEC

In CEC, the EOF drives the mobile phase through the capillary. The generation of EOF in CEC is similar to what was described in Section 2.3, except now the packing material and the capillary wall both contribute to the total flow

Since the EOF depends on the free silanols in the column packing, the type of packing is very important. The porosity of the media plays a crucial role here as well. Nonporous packings yield a higher EOF than porous packings of the same particle diameter (1). Table 7.2 lists a variety of chromatographic media along with values for the generated EOF

Table 7.2 Electroosmotic Mobilities on Some Chromatographic Materials

Stationary Phase Material Electroosmotic Mobility (x 10"" cmWs)

Setl

CEC Hypersil Cis 2.26

ODSHypersil 1.47

BDS-ODS Hypersil 0.99

Sperisorb ODS 1 2.26

Sperisorb ODS II 1.79

Set 2

Nucleosil 5 Cig 1-56

LiChromospher RP-18 1.45

Sperisorb Dial 0.80

Zorbax BP-ODS 0.68

Sperisorb 55 ODS2 0.50

Hypersil ODS 0.14

Partisil 5 ODS3 <0.01

Purospher RP-18 <0.01

Data from (18), which consisted of data from (19, 20). Conditions: thiourea used as unretained marker; set 1: 80:20 acetonitrile: Tris-HCl, 50 mM, pH 8, 20°C; set 2: 70:30 acetonitrile: 3-cyclohexylamino-2-hydroxy-l-propane-sulfonic, 25 mM.

3 0 0 Chapter 7 Capillary Electrochromatography

The electroosmotic velocity is described by the Smoluchowsi model (1), where

riL

Here e^ is the permittivity of vacuum, £^ is the dielectric constant of the BGE, ^ is the zeta potential, and 7] is the viscosity These factors are identical to the controlling forces in CZE. Changes in temperature, pH, viscosity organic solvents, and ionic strength all affect the EOF This model considers the column in CEC as a bundle of capillaries. In this regard, the magnitude of the EOF would not depend on the particle size, as long "double-layer overlap"^ does not occur.

A. EFFECT OF VOLTAGE

At field strengths above 100 V/cm, the EOF exhibits a positive deviation from linearity This may be due to Joule heating effects, but it also could result from polarization of the double layer. In this case, the particles become more conductive than the surrounding electrolyte (1, 22). In any event, the higher than expected EOF might be advantageous in speeding the separation, although reproducibility might be lost.

B. EFFECT OF BUFFER CONCENTRATION

The addition of a buffer to the BGE is necessary to adjust pH and maintain electrical conductivity, although there has been a report on CEC in salt-free media (23). The zeta potential and, thus, the EOF are inversely proportional to the square root of the buffer concentration.

C. EFFECT OF ORGANIC

MODIFIER CONCENTRATION

In CZE, the organic modifier concentration can alter the EOF based on changes of the viscosity of the BGE. Acetonitrile effects only modest changes in EOF, whereas linear alcohols reduce the EOF as the concentration is increased. In CEC, depending on the experimental conditions, the EOF may increase or decrease as the acetonitrile concentration is raised from 0% to 60% (1).

iThis is defined as the overlap of the electrical double layer that gives rise to the EOF. It results in a reduction in EO¥, but this is not observed until the particle size of the packing is well below 1 /im (21).

7.4 Efficiency of CEC 3 0 1

7.4 EFFICIENCY OF CEC

In open tubular CEC, two processes contribute to band broadening in capillary chromatography: diffusion and mass transfer in the mobile phase. These contributions can be expressed via the Van Deemter equation (24)^

H = 2 ^ + C ^ — , (7.3)

where H = height equivalent to a theoretical plate; V = mean linear velocity; Djn = solute diffusion coefficient in the mobile phase; r = the capillary radius; and Cjn = the coefficient of resistance to mass transfer for a solute in the mobile phase.

The first term in the equation describes the time-related impact of solute diffusion. This term is equivalent for HPLC and CEC. Since there is no packing material in open tubular CEC, the "so-called A" term (see below) of the Van Deemter equation is absent. Also absent from this equation are terms describing mass transport in the stationary phase. This is an unimportant source of band broadening both in HPLC and CEC.

Differences between CEC and HPLC are manifested in the mass transfer term in the mobile phase. This function is affected by the cross-sectional flow profile. In pressure-driven systems, the height of the theoretical plate (H) is given by

H = l ± ^ ^ : ± l l ^ . (7.4) 96(1 + k'f

In voltage-driven systems,

H - ^ ^ . (7.3) 16(1 + k'f

These equations can be used to assess the impact of laminar flow in the pressure-driven system versus plug flow in the electrodriven case. From this simple analysis, the concepts can be translated to the packed capillary

Calculated values based on these equations along with their ratio are plotted versus k' in Figure 7.4. At k' values of greater than 1, the ratio Hpressure/ voitage has a value of 2. This indicates the electrodriven system is only twice as efficient as the pressure-driven system. When k' is less than 0.5, the electrodriven system shows a huge advantage. In this case, the system is functioning close to zero retention, and the efficiencies become more like CZE. Here, the mass transfer contributions

^The letter v is used to define velocity instead of fi. This is to avoid confusion between elec-trophoretic mobility and velocity.


H 0.06

0.04

0.02

Press Voltage

FIGURE 7.4 Comparison of the height of the theoretical plate for pressure- and electropumped systems versus k'. The graph is based on calculations from Eqs. (7.4) and (7.5).

toward band broadening become insignificant, and the laminar flow profile of the pressure pumped system becomes the most significant source of band broadening.

In the packed capillary, the Van Deemter equation becomes (25)

H = 2Adp 4- ^yD^

30

r k' ] 2 f J2 \

(7.6)

where A = the tortuosity factor; dp = the particle diameter; y = the obstructive factor for diffusion; v = the migration velocity, and D^^^ = the diffusion in the stagnant mobile phase. The first term of this equation (A term) represents the contribution of flow variation in the column packing. This is otherwise known as eddy diffusion.

The further apart adjacent particles are from each other, the larger the inter-particle flow velocity profile. This is illustrated in Figure 7.5. When the elec-trodriven system is considered, the flow profiles are now uniform despite differences in the interparticle distances. Analyzing the relevant mathematics and Van Deemter plots, we find that the reduction of eddy diffusion appears responsible for the improved efficiency of CEC over HPLC (25, 26).

Rigorous calculations support the reduction of parabolic flow in CEC from that in HPLC as the factor that improves efficiency (25, 26). Experimental data given in Figure 7.6 further support the modest improvements in efficiency for electropumped systems over pressure-pumped systems under uniform conditions. An expansion of the fluorene peak is shown in Figure 7.7. The improvements in efficiency are significant but not dramatic.

Not considered in this analysis is the upper limit for pumping pressure in HPLC. This limits the length of the column when small particles are employed. In CEC, there is no pressure drop and, thus, no limit to the length of the capillary. The impact of column length and particle diameter is shown in Table 7.3.

7.5 Operating Characteristics of Packed CEC 3 0 3

Pmssurt driva

Electroendosmotlc drive »»i .Mn»,»^

particle —** ^P™

channel

ed >

(E

velocity profll©

M 3

FIGURE 7.5 Diagram of eddy diffusion and flow velocity profiles in HPLC and CEC. Courtesy of Hewlett-Packard Corporation.

Using 5-|im particles and a 50-cm capillary, the CEC separation is approximately twice as efficient as capillary HPLC, in agreement with the theoretical prediction.

When the particle size is reduced to 3 |im, the HPLC column length can be no longer than 25 cm because of the 400-bar pressure limit. This limits the plate count to 45,000 by HPLC, whereas the CEC separation using a 50-cm capillary yields 170,000 theoretical plates. When L5-|im material is used, the 50-cm CEC capillary yields 250,000 plates, compared with 30,000 plates on a 10-cm HPLC capillary

7.5 OPERATING CHARACTERISTICS OF PACKED CEC

A. PREPARATION OF CEC CAPILLARIES

While most users will purchase prepacked capillaries, the steps in column preparation are worth considering (27). These are illustrated in Figure 7.8 on p. 306 and described in the following:

1. Prepare a temporary outlit frit by sintering silica gel particles with a microflame torch.

304 chapter 7 Capillary Electrochromatography

PRESSURE

yjL O 10

ELECTRO 20 30

lyL 10 20 30

TIME (min) 40

FIGURE 7.6 Separation of polycyclic aromatic hydrocarbons on a drawn capillary packed with 3-|Lim Hypersil particles and derivatized in situ with octadecylsilane. Capillary: 90 cm (pressure), 80 cm (electro) x 30 |J.m i.d.; pressure (upper): 25 bar; voltage (lower): 320 V/cm; detection: fluorescence. Order of elution: naphthalene, 2-methylanthracene, fluorene, phenanthrene, anthracene, pyrene, and 9-methylanthracene. Reprinted with permission from Chromatographia, 32, 317 (1991) copyright © 1991 Vieweg.

2. Slurry pack the capillary with packing media dispersed in methanol at 350 bar for 3 h.

3. Prepare the permanent outlet frit using a thermal wire stripper. 4. Unpack the capillary by pumping from both ends. 5. Prepare the detector window. 6. Repack the capillary as in step 2. 7. Prepare the inlet frit by gently sintering the particles at the end of the

capillary.

While there are certainly many variations of this technique, the packing of these capillaries is an art form, and there is a high reject rate. The frit in particular is prone to problems. Frits must retain the chromatographic packing yet be sufficiently porous to allow the passage of solvent. Details of frit production have been described, along with recommendations (28). Frits prepared from

7.5 Operating Characteristics of Packed CEC 305

PRESSURE / / \ \ ELECTRICiUXY OraVEN \ / / \ \ DRIVEN

612 600 588 DISTANCE (mm)

FIGURE 7.7 Expansion of the fluorene peak from Figure 7.6. Outer curve: pressure-driven chro-matogram; inner curve: electrically driven chromatogram. Reprinted with permission from Chro-matographia, 32, 317 (1991) copyright © 1991 Vieweg.

pure spherical silica gel appear best. Since production of frits by sintering destroys the polyimide, the capillary must be carefully handled.

Packed capillaries are available from many sources, including Hewlett-Packard, Hypersil, Micro-Tech Scientific, Capital HPLC Ltd., Phase Separations, and Unimicro Technologies, Inc.

B. COLUMN EQUILIBRATION

The capillary must be flushed with mobile phase prior to use. This can be done with an external HPLC pump, a manual syringe pump,^ or the instrument internal pressure, particularly if a high-pressure mode is available. It may take several hours to totally condition a capillary. When all air bubbles have left the capillary, it is best to further condition the capillary at low voltage.

Alternatively, the capillary may be conditioned by pressurizing the inlet to 10 bar, running a voltage gradient to 25 kV over 30 min, and holding the voltage at 25 kV for an additional 30 min (29).

^Procedure given in Unimicro Technologies operating instructions.

Table 7.3 Achievable Plate Numbers in Capillary HPLC and CEC

Particle Size (pm)

5

3

1.5

Capillary

Length (cm)

50

25

10

HPLC

Plates/Column

55,000

45,000

30,000

Length (cm)

50

50

50

CEC

Plates/Column

115,000

170,000

250,000


Temporary outlet frit

4

5 3

Outlet frit inlet frit Detector window

FIGURE 7.8 Illustration of the processes for frit production and packing of a CEC capillary.

C. INJECTION

Electrokinetic injection is usually employed in CEC. The mobility discrimination often seen in CZE is less significant in CEC, since the EOF is the driving force for injection. Hydrodynamic injection can also be employed if a high-pressure injection mode is available. Since only nanoliters of material must enter the capillary, 8 bar of pressure for 30 s provides a sufficient injection (30).

Sample loading is improved by injecting the sample dissolved in a solvent containing more water than the mobile phase (31). In this case, the solutes are retained at the head of the packed capillary and then are eluted by the mobile phase. This increases the loading capacity of the system to the nanogram range.

D. CAPILLARY AND MOBILE-PHASE SELECTION

Start this process by selecting packings and mobile phases similar to those that have been successful in HPLC. The data given in Table 7.2 should be useful in

7.5 Operating Characteristics of Packed CEC 307

ensuring that a packing material with good EOF characteristics is selected. In addition to C g material, octyl, phenyl, cyano, and amino reversed-phase materials are available. If specialty phases are required and a packing apparatus is available, a purchased HPLC column will provide a lifetime supply of packing material.

Selection of particle sizes of 3 |im or less will yield the most efficient separations. Using reversed-phase material, select a pH no less that 2.5 (to maintain EOF) nor greater than 9 (to prevent silica dissolution). At pH 2.5, the EOF is 5 to 10 times lower than at pH 7.

Use buffers such as Tris, phosphate, borate, and acetate at concentrations from 5 to 25 mM to adjust pH and maintain conductivity. The buffer concentration should be optimized, as chromatographic efficiency often improves at higher (25 mM) concentrations. If the buffer concentration is too high. Joule heating may adversely affect the efficiency of the separation.

To avoid the pH dependency of the EOF, cation- or anion-exchange packings can be used. A propylsulfonic acid, either by itself or mixed with CIQ material (mixed-mode packing), can provide sufficient EOF, even at acidic pH (30).

Acetonitrile is best as a mobile-phase modifier, because unlike methanol, it does not lower the EOF That is always an important consideration when selecting the appropriate modifier. Typical acetonitrile concentrations are 20-80%, depending on the application. Acetonitrile also is more optically transparent than methanol, which can improve detectability Before selecting a solvent other than methanol or acetonitrile, ensure that the material is compatible with the instrument, vials, and vial closures.

Methanol can also be used as a mobile-phase modifier when acetonitrile is not appropriate. When using low concentrations of methanol or acetonitrile, the wetting characteristics of the BGE are poor. This can lead to bubble formation, which interrupts the electric circuit. Pressurizing the system with 8-10 bar over both the inlet and outlet vials will minimize bubble formation. The use of low-conductivity buffers such as Tris and MES has also been reported to help suppress bubble formation (32).

Another way of suppressing bubble formation is to add small amounts of surfactants to the system. When using submicellar concentrations of SDS (1-5 mM), bubble formation is suppressed and the EOF may be stabilized (33). In this case, 20% methanol was the modifier. Such a low concentration of modifier permits the SDS to bind to the packing material and modify the stationary phase. At higher modifier concentrations, such binding does not occur, since the organic solvent effectively keeps the SDS off the packing material.

Yet another way to suppress bubble formation is to use supplementary |I-HPLC pressurized flow. The application of a voltage to a |I-HPLC separation clearly improves the chromatographic efficiency Another stated advantage is that the separation does not entirely rely on the packing to produce the EOF (34, 35). This approach has been named pressure-driven CEC or electro-HPLC. Since the


capillary is coupled to an LC system, the formation of gradients is simpler as well. However, it should be noted that one of the previously cited papers showed superior separations of oligonucleotides using MECC (34).

While instruments employing gradient elution are just being introduced, a step gradient can be designed with almost any commercial instrument (36). The separation begins with the weaker mobile phase, and after a period of time, the voltage is removed and the inlet and outlet vials replaced with the stronger eluent. After completion of the run, fresh weak solvent is used to reequilibrate the capillary

E. VOLTAGE

Most instruments function at a maximum voltage of 30 kV Higher voltages will result in rapid analysis, provided Joule heating is insignificant. Use of a 20-cm capillary with 55-kV applied voltage provides an isocratic separation of 16 poly-cyclic aromatic hydrocarbons in 2 min (37).

F. DETECTION

Postfrit detection is always employed when using UV absorbance detection, since the capillary packing is opaque. Extended path length capillaries can also be used to increase the sensitivity of detection (29).

When using LIF detection, on-capillary detection is possible (37). This approach yields the highest chromatographic efficiency (700,000 plates/meter) and indicates that postfrit detection results in band broadening.

The absence of micelles in CEC is advantageous with regard to mass spectrometry (31, 38, 39). Typical mobile phases contain acetonitrile, ammonium acetate (31), or trifluoroacetic (40). A sheath flow of a few microliters per minute is frequently used to provide a stable electrospray, since the flow rate through the column is quite small. Through the use of nanoelectrospray, the sheath fluid becomes unnecessary.

Moving closer to cutting edge technology, open tubular CEC has been interfaced to a time-of-flight instrument via the nanoelectrospray. The advantage of the time-of-flight instrument is that it is nonscanning. In conjunction with an ion trap, which enriches the ions, limits of detection for peptides reach 10" M (41). CEC has also been interfaced to a nuclear magnetic resonance spectrometer (42).

G. TEMPERATURE

As in all other forms of HPCE, temperature control is particularly important. When increasing the capillary temperature from 20°C to 60°C, the retention time %RSD was lowered from 7.3% to 4.3%, and the analysis time was short-


ened by 45% (43). To keep the current low and aid in the suppression of bubble formation, low-temperature operation at 15°C has been recommended (30).

7.6 APPLICATIONS

A representative selection of applications and chromatographic conditions is given in Table 7.4. Some other applications of note are described in this section.

The open tubular separation of sulfonic acids (Figure 7.9, p. 312) on a 10-|Xm-i.d. capillary coated with 0.9% PS-264 or 10% OV-17 employs an ion-pairing reagent, tetrabutylammonium hydroxide, as a mobile-phase modifier (49). The use of a narrow-bore capillary improved the mass transfer problem in accordance with the second term of Eq. (7.3). Improvements over CZE and pressure-driven LC separations were demonstrated with this ion-pair CEC system. Thus, the chemistry employed in conventional HPLC separations can be used in CEC as well.

Separations of the drug Isradipin and its by-products (Figure 7.10, on p. 313) (45) present a good example of a separation with small V values (0.17-0.90). The large number of plates per unit of time is consistent with the theory, which predicts optimal efficiency at small fe' values. In this early work, the retention time precision ranged from 1.6% to 2.2% (run/run) and 9% (capillary/capillary). Production of the narrow-bore packed capillaries proved problematic in this early work and still does.

High-speed separations by CEC are best accomplished by maximizing the electric field strength. Since most instruments are incapable of producing greater that 30 ky it is necessary to use short capillaries to provide the high electric field. This is effective when the separations are not too complex. Figure 7.11, on page 314 shows the separation of some aromatic hydrocarbons using 1.5-|lm nonporous ODS particles packed in a 6.5 cm (10 cm total length) X 100 |Xm capillary (37). With the voltage set at 28 kV, the field strength is 2800 V/cm. Above 28 ky arcing between the capillary and the capillary holder was observed. This becomes another limiting factor when using high voltage, since the grounding and shielding within the instrument become most critical. On commercial instrumentation, it is usually necessary to use the "short end" of the capillary to produce such a short capillary length (52).

Another factor that permitted operation at such a high field strength was the low conductivity of the BGE. With 2 mM Tris in 70% acetonitrile, the current was 6.7 |LiA at 30 kV. An Ohm's law plot showed a 20% deviation from linearity at 20 ky so that this homemade system could use better heat dissipation.

Combined with on-capillary LIE detection to minimize dispersion from the frits, the separation time for this simple mixture is less than 5 s. With a peak width of 0.2 s, it is necessary to set the detector time constant to 0.02-0.03 s to minimize band broadening, and the injection size must be kept quite small. Like other forms of HPCE, whenever the peak widths become narrow, all instrumental parameters must be carefully controlled to maintain the inherent efficiency of the separation.

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FIGURE 7.9 Electrochromatography of naphthalene sulfonic acids. Capillary: 50 cm x 10 |xm i.d. PS-264 coated; buffer: 10 mM phosphate, pH 7, 1.25 mM tetrabutylammonium hydroxide; voltage: -21 kV; detection: laser fluorescence. Key: 4A1N, 4-amino-l-naphthalene sulfonic acid; 2A1N, 2-amino-l-naphthalene sulfonic acid; 8A2N, 8-amino-2-naphthalene sulfonic acid; 5A2N, 5-amino-2-naphthalene sulfonic acid; 1H4N, l-naphthol-4-sulfonic acid. Reprinted with permission from J. Chromatogr., 557, 125 (1991) copyright © 1991 Elsevier Science Publishers.

CEC is beginning to have an impact in the field of chiral recognition (35, 53, 54). As an example, a separation of thalidomide enantiomers on an immobilized vancomycin chiral stationary phase is shown in Figure 7.12 (55). Vancomycin is a powerful chiral selector. The problem with its use in CZE is its high-UV absorbance. This often requires that special techniques such as partial capillary filling be used (Section 4.9F). In CEC, no such problems occur, since the antibiotic is bound to the capillary wall. While Figure 7.12 shows a nonaqueous separation in the "polar-organic mode," reversed-phase separations work as well but are 30% less efficient.

The production of a bound vancomycin column began with packing a diol silica into a 100-|im capillary. The diol was oxidized with sodium periodate. Vancomycin was attached by reductive amination (in 50 mM phosphate, pH 7) with sodium cyanoborohydride followed by reduction of free aldehyde, again with sodium cyanoborohydride (in 50 mM phosphate, pH 3) on the packing material. It is these types of creative approaches that will expand the scope and future applications of CEC.

During studies employing cation-exchange packings, some unusual results were noted. For basic compounds, this packing yielded "staggering efficien-

7.7 CEC Microfluidic Devices 313

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FIGURE 7.10 Capillary electroosmotic chromatography of Isradipin and its by-products. Capillary: 14.3 cm X 50 lim i.d. packed with Hypersil ODS (3 \x.m)\ mobile phase: 2 mM sodium tetraborate (pH 8.7)-80% acetonitrile; voltage: 30 kV; injection: 4 s at 1.5 kV; detection: UV, wavelength not specified. Reprinted with permission from J. Chromatogr., 593, 313 (1992) copyright © 1992 Elsevier Science Publishers.

cies," but explanations were not forthcoming. This effect was first observed by Smith and Evans (56). The possibihty of some form of isotachophoresis (Section 8.61) was considered but never proved. In any event, milUons of theoretical plates were observed. Similar results were observed by Moffatt et ah (57) for partially ionized anionic or neutral compounds using a Cis- Their explanation of the effect is nonequilibrium conditions caused by pulses of strong or weak solvent that arise from the sample. The increased efficiency occurs when the migration time of the solute is matched to the elution time of the sample-induced discontinuity. As shown in Figure 7.13, focusing does not occur when the solute is dissolved in the mobile phase. The peak width of the neutral marker, thiourea, is the same in both instances. Good migration time and peak area reproducibility were found, indicating that the focusing effect can be considered for analytical applications where high sensitivity and resolution are required (57).

7.7 CEC MICROFLUIDIC DEVICES

With the advent of micromachining to produce chemical analysis systems, it becomes possible to perform CEC on these chip-based systems (58, 59). The design of one such column is shown in Figure 7.14 (59). The capillary inlet is


2 3 4 5 6 RETENTION TIME (SECONDS)

FIGURE 7.11 High-speed separation of polycyclic aromatic hydrocarbons on a short capillary at high field strength. Capillary: 6.5 cm (10 cm total length) x 100 |xm; packing: 1.5-|xm non-porous CDS material; mobile phase: 70% acetonitrile, 2 mM Tris; voltage: 28 kV; injection: 5 kV for 2 s; detection: LIF, doubled argon-ion laser at 257 nm, emission collected between 280 and 600 nm. Reprinted with permission from Anal. Chem., 70, 4787 (1998) copyright © 1998 Am. Chem. Soc.

mAU 100

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FIGURE 7.12 Separation of thahdomide enantiomers on an in situ immobilized vancomycin chi-ral stationary phase. Capillary: 26 cm x 100 mm; packing: diol silica with immobilized vancomycin; mobile phase: 80% methanol, 20% acetonitrile, 0.2 parts acetic acid, 0.2 parts triethylamine; voltage: 20 kV; temperature: 15°C; detection: UV, 260 nm; pressurization: 10 bar over inlet and outlet. Courtesy of Paul Owens, Astra Hassle AB, Sweden.

7.7 CEC Microfluidic Devices 315

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FIGURE 7.13 High-efficiency CEC separation caused by an unusual focusing effect. Capillary: 33 cm (24.5 cm to detector) x 50 mm; packing: 3-mm Hypersil Cig; mobile phase: 70:30 [5 mM Tris (pH 8.6):acetonitrile]; injection: 5 kV for 5 s; voltage: 30 kV; temperature: 30°C; detection: Uy 214 nm; inlet and outlet pressurized with nitrogen at 10-12 bar; solute: 6-(hydroxymethyl)-2-(methylamino)-5-methylpyrimidin-4-ol, 2.1 mg/mL. Solute dissolved in (A) water; (B) mobile phase. Reprinted with permission from Anal. Chem., 71, 1119 (1999) copyright © 1999 Am. Chem. Soc.

split several times to allow the sample to be dispersed over the functionalized surface of the etched column. A similar structure occurs at the outlet to allow recombination of the channels for detection. It appears likely that there will be a role for CEC in the forthcoming revolution in miniaturized analytical systems.


COMOSS f l

Collocated Monolith Collector

-8d-

16d

-8d-

16d

Coupling Channels

-32d- 1

No. of Channels

FIGURE 7.14 Configuration of an inlet splitter to disperse sample and mobile phase over a micro-fabricated chromatographic column. Reprinted with permission from Ana/. Chem., 70, 3790 (1998) copyright © 1998 Am. Chem. Soc.

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9. Rathore, A. S., Horvath, C. Effect of a Predetection Open Segment in the Column on Speed and Selectivity by Capillary Electrochromatography. Anal. Chem., 1998; 70:3271.

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13.Peters, E. C, Petro, M., Svec, F., Frechet, J. M. J. Molded Rigid Polymer Monoliths as Separation Media for Capillary Electrochromatography 1. Fine Control of Porous Properties and Surface Chemistry. Anal. Chem., 1998; 70:2288.

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17. Fujimoto, C. Charged Polyacrylamide Gels for Capillary Electrochromatographic Separations of Uncharged, Low Molecular Weight Compounds. Anal. Chem., 1995; 67:2050.

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21. Knox, J. H., Grant, 1. H. Miniaturisation in Pressure and Electroendosmotically Driven Liquid Chromatography: Some Theoretical Considerations. Chromatographia, 1987; 24:135.

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25. Knox, J. H. Thermal Effects and Band Spreading in Capillary Electro-separation. Chro-matographia, 1988; 26:329.

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33.Bailey, C. G., Yan, C. Separation of Explosives Using Capillary Electrochromatography. Anal. Chem., 1998; 70:3275.


34.Behnke, B., Bayer, E. Pressurized Gradient Electro-High-Performance Chromatography. J. Chro-matogr.. A, 1994; 680:93.

35.Deng, Y., Zhang, J., Tsuda, T., Yu, P H., Boulton, A. A., Cassidy R. M. Modeling and Optimization of Enantioseparation by Capillary Electrochromatography. Anal Chem., 1998; 70:4586.

36.Euerby, M. R., Gilligan, D. Step-Gradient Capillary Electrochromatography. Analyst, 1997; 122:1087.

37. Dadoo, R., Zare, R. N., Yan, C, Anex, D. S. Advances in Capillary Electrochromatography: Rapid and High-Efficiency Separations of PAH's. Anal Chem., 1998; 70:4787.

38. Lane, S., Boughtflower, R., Paterson, C, Morris, M. Evaluation of a New Capillary Electrochro-matography/Mass Spectrometry Interface Using Short Columns and High Field Strengths for Rapid and Efficient Analyses. Rapid Commun. Mass Spectrom., 1996; 10:733.

39. Lord, G. A., Gordon, D. B., Tetler, L. W, Carr, C. M. Electrochromatography-Electrospray Mass Spectrometry of Textile Dyes. J. Chromatogr., A, 1995; 700:27.

40. Wu, J.-T., Hunag, P, Li, M. X., Lubman, D. S. Protein Digest Analysis by Pressurized Capillary Electrochromatography Using an Ion Trap Storage/Reflectron Time-of-Flight Detector. Anal Chem., 1997; 69:2908.

41. Wu, J.-T., Huang, P., Li, M. X., Qian, M. G., Lubman, D. M. Open-Tubular Capillary Electrochromatography with an On-Line Ion Trap Storage/Reflection Time-of-Flight Mass Detector for Ultrafast Peptide Mixture Analysis. Anal Chem., 1997; 69:320.

42. Pusecker, K., Schewitz, J., Gfrorer, P, Tseng, L.-H., Albert, K., Bayer, E. Online Coupling of Capillary Electrochromatography, Capillary Electrophoresis, and Capillary HPLC with Nuclear Magnetic Resonance Spectroscopy. Anal Chem., 1998; 70:3280.

43. Djordjevic, N. M., Fowler, P W. J., Houdiere, E, Lerch, G. Retention of Neutral Solutes by Capillary Electrochromatography J. Liq. Chromatogr. Related Technol, 1998; 21:2219.

44. Yan, C, Schaufelberger, D., Emi, E Electrochromatography and Micro High-Performance Chromatography with 320 |Xm I.D. Packed Columns. J. Chromatogr, A, 1994; 670:15.

45.Yamamoto, H., Baumann, J., Erni, F. Electrokinetic Reversed-Phase Chromatography with Packed Capillaries. J. Chromatogr, 1992; 593:313.

46. Rebscher, H., Pyell, U. A Method for the Experimental Determination of Contributions to Band-broadening in Electrochromatography with Packed Capillaries. Chromatographia, 1994; 38:723.

47. Li, S., Lloyd, D. K. Packed-Capillary Electrochromatographic Separation of the Enantiomers of Neutral and Anionic Components Using j3-Cyclodextrin as a Chiral Selector Effect of Operating Parameters and Comparison with Free-Solution Capillary Electrophoresis. J. Chromatogr, A, 1994; 666:321.

48. Smith, N. W., Evans, M. B. The Analysis of Pharmaceutical Compounds Using Electrochromatography. Chromatographia, 1994; 38:649.

49.Pfeffer, W. D., Yeung, A. S. Electroosmotically Driven Electrochromatography of Anions Having Similar Electrophoretic Mobihties by Ion Pairing. J. Chromatogr, 1991; 557:125.

50.Dittmann, M. M., Rozing, G. P, Ross, G., Adan, T., Unger, K. K. Advances in Capillary Electrochromatography. J. Capillary Electrophor, 1997; 4:201.

51. Yan, C, Dadoo, R., Jui, Z., Zare, R. N., Rakestraw, D.J. Capillary Electrochromatography: Analysis of Polycyclic Aromatic Hydrocarbons. Anal. Chem., 1995; 67:2026.

52.Euerby, M. R., Johnson, C. M., Cikalo, M., Bartle, K. D. "Short-End Injection" Rapid Analysis Capillary Electrochromatography. Chromatographia, 1998; 47:135.

53.Schweitz, L., Andersson, L. I., Nilsson, S. Capillary Electrochromatography with Molecular Imprint-Based Selectivity for Enantiomer Separation of Local Anesthetics. J. Chromatogr, A, 1997; 792:401.

54. Peters, E. C, Lewandowski, K., Petro, M., Svec, F, Frechet,J. M.J. Chiral Electrochromatography with a "Molded" Rigid Monolithic Capillary Column. Anal. Commun., 1998; 35:83.

55. Wikstrom, H., Svensson, L. A., Owens, P K. Vancomycin Chiral Stationary Phase for Capillary Electrochromatography, in HPCE'99. 1999.

References 3 1 9

56. Smith, N. W., Evans, M. B. The Efficient Analysis of Neutral and Highly Polar Pharmaceutical Compounds Using Reversed-Phase and Ion-Exchange Electrochromatography. Chromato-graphia, 1995; 41:197.

57. Moffatt, E, Cooper, P A., Jessop, K. M. Capillary Electrochromatography. Abnormally High Efficiencies for Neutral-Anionic Compounds under Reversed-Phase Conditions. Anal Chem., 1999; 71:1119.

58.Kutter, J. P, Jacobsen, S. C, Matsubara, N., Ramsey, J. M. Solvent-Programmed Microchip Open-Channel Electrochromatography. Anal. Chem., 1998; 70:3291.

59. He, B., Niall, T., Regnier, E Fabrication of Nanocolumns for Liquid Chromatography. Anal. Chem., 1998; 70:3790.

CHAPTER 8 Injection

8.1 Volumetric Constraints on Injection Size 8.2 Performing an Injection and a Run 8.3 Injection Techniques 8.4 Short-End Injection 8.5 Injection Artifacts: Problems and Solutions 8.6 Stacking and Trace Enrichment

References

8.1 VOLUMETRIC CONSTRAINTS ON INJECTION SIZE

All capillary separation techniques including HPCE have certain constraints on the amount of material that can be injected. Injection in HPCE is designed to allow introduction of sufficient material into the capillary and minimize the extracapillary variance from the process itself. The volumetric problem is expressed in Table 8.1. Because the entire internal volume of a 50 cm x 50 |im i.d. capillary is only 981 nL, the injection volume must be kept quite small.

In Section 2.14, the concept of additivity of variances was introduced (1-3). This will now be used to calculate the impact of injection on the theoretical plate count.

The contribution to variance from a plug injection is (1, 2)

2

12 < = ^ , (8-1)

Table 8.1 Internal Volume versus

Capillary i.d. (vim)

Volume/mm (nL)

Total volume (nL)

10

0.0785

39.3

Diameter for a

25

0.491

245

50-cm Capillary

50

1.96

981

100

7.85

3,925

200

3L4

15,700

321

322 Chapter 8 Injection

where / is the length of the injection plug. To model the increase in band broadening from the injection process, the diffusion-limiting case must be considered via the Einstein equation:

aU=2D^t. (8.2)

Since the squares of the variances are additive, the contributions to band broadening from injection and diffusion can be inserted into the theoretical plate equation:

(8.3)

For a 50-cm capillary and a solute migration time of 600 s, the impact of the injection size for a small molecule (D^ = 10"^ cmVs) and a large molecule (D^ - 10"^ cmVs) is shown in Figure 8.1. As can be seen from the figure, injection of 1% of the capillary volume with sample causes a 92% loss of efficiency for a large molecule and an 8% loss of efficiency for the prototypical small molecule. As discussed in Section 2.14, the more efficient the separation, the harder it is to maintain that efficiency.

The same can be seen in Table 8.2, where calculations have been made to predict the allowable injection sizes that provide 5% and 10% increase in peak

T H E O R E T 1 C A L

P L A T E S

1000000

100000

10000 0.1 0.2 0.3 0.4 0.5 0.6 0.7

INJECTION SIZE (cm)

0.8 0.9

FIGURE 8.1 Impact of injection zone length on the number of theoretical plates for a small molecule, 0 (Djn = 10"^ cmVs), and a large molecule, A {D^ = 10~^ cmVs), as solved by Eq. (8.3). Conditions: capillary length: 50 cm to detector; migration time: 600 s.

8.2 Performing an Injection and a Run 323

Table 8.2 Calculated Maximum Injection Length and Volume for a 50 cm X 50 pim i.d. Capillary

Number of Theoretical Plates (N) Peak Width (mm),

5% increase

linj (mm)

Vinj (nL)

10% increase

/inj (mm)

Vinj (nL)

tm-= 10 min 100,000

4.47

1.24

2.43

1.77

3.48

500,000 2.00

0.56

1.09

0.79

1.55

1,000,000 1.41

0.39

0.77

0.56

1.10

Data from reference (2). Peak widths at half height calculated from N = 5.5^(,t^^/w'^y ).

width for separations yielding 100,000, 500,000, and 1,000,000 theoretical plates (2). To maintain 95% efficiency for a 1,000,000-plate separation, an injection plug of 0.07% of the capillary length is required. For a 100,000-plate separation, a 0.24% injection can be tolerated.

Both of these models assume that the injection electrolyte is identical to the running electrolyte. Through the use of a low-conductivity injection solution relative to the BGE, it is possible to obtain substantial compression of the injection zone. Introduced in Chapter 2, this process is known stacking. This family of related processes will be described in detail later in this chapter.

In any event, the problem of injection is one of the driving forces behind micromachined systems. Using "pinched injection," it is possible to inject minute amounts of sample (4) without suffering the effects of "ubiquitous injection" (see Section 8.5). This allows shortened capillaries to be employed in complex separations, such as the separation of DNA-sequencing reaction products. Through the use of LIF detection, sensitivity is not a problem despite the small injection size.

To approach the optimal efficiency of HPCE, injection must be optimized. Through the use of dilute aqueous samples and short electrokinetic injections, separations providing millions of theoretical plates are possible (5). As is often the case, efficiency will often be sacrificed to optimize sensitivity.

8.2 PERFORMING AN INJECTION AND A RUN

The steps involved in injecting a sample into a capillary and performing a run are as follows:

1. Equilibrate the capillary in run buffer for 1-5 min. 2. Transfer the sample vial to the capillary-electrode assembly.

3 2 4 Chapter 8 Injection

3. Immediately inject the sample via electrokinetic or hydrodynamic injec-t ion(l-30s) .

4. If necessary to eliminate carryover, designate a wash station to rinse the outside walls of the capillary.

5. Return the capillary-electrode assembly to the sample-side run buffer. 6. Promptly engage the high voltage using a 15-s voltage ramp to maximum

voltage, and allow the separation to proceed. 7. If necessary, run a wash step with 0.1 N or 1 N sodium hydroxide or

phosphoric acid.^ 8. Return to step 1 for the next sample.

It is important to proceed with the run as soon as the sample is loaded into the capillary Otherwise, solute diffusion will broaden the injection zone (7). In addition, solute ions will diffuse into the BGE, and conversely, BGE ions will diffuse into the capillary.

8.3 INJECTION TECHNIQUES

Injection probably contributes the greatest analytical error in a well-controlled method. In HPLC, a fixed loop precisely defines the amount of material injected into the system. In HPCE, this is not the case, as will be seen in the following discussion.

In HPLC, injection is conducted with a flowing mobile phase. A valve is used to isolate the loop, allowing injection to be made at atmospheric pressure. The valve is then activated to sweep the sample onto the chromatographic column. For HPCE, injection is made with the voltage off. There have been numerous reports describing schemes employing loops (8) or that are designed to permit injection with the voltage on (9, 10), but none are found on commercial instruments.

After injection, the voltage is ramped up to the operating potential. Both of these processes contribute to analytical error, which can easily be as high as 1% in well-controlled methods. The use of internal standards is required to eliminate this source of error.

The voltage ramp is required to retain the sample within the capillary. If the full voltage is applied at once, the BGE heats up and expands. If the rate of fluid expansion is greater than the rate of solute migration in the capillary, some of the sample may backwash into the inlet reservoir (11). The voltage ramp prevents this from occurring. An alternative method is to inject a small plug of BGE

iThe wash step can include 100 mM SDS if proteins are endogenous components of the sample (6).

8.3 Injection Techniques 325

to move the sample into the capillary. This method is not preferred, since hydro-dynamic flow causes a small amount of band broadening.

There are two forms of injection used in HPCE—hydrodynamic injection and electrokinetic injection. Hydrodynamic injection is simple to employ and usually guarantees that the proper amount of sample enters the capillary. With electrokinetic injection, the conductivity of the sample relative to the BGE influences the number of ions entering the capillary. This parameter must be carefully controlled if quantitative results are required. The advantage of electrokinetic injection is that extreme trace enrichment is possible. These two modes of injection are described in the following subsections.

A. HYDRODYNAMIC INJECTION

Hydrodynamic injection is accomplished in one of four ways:

1. By elevating the capillary at the sample (inlet) end, permitting sample introduction by siphoning

2. By applying pressure on the individual sample vial 3. By applying vacuum on the detector-side buffer reservoir 4. Injecting by syringe and employing a splitter to reduce the volume intro

duced into the capillary

Until the arrival of commercial instrumentation, nearly all hydrodynamic injections were performed using method 1. When performed manually, as was frequently the case, injection precision was often quite poor. Instruments manufactured by Waters and Dionix used the technique by automatically raising the inlet side using a pedestal mechanism. Most other commercial systems employ methods 2 or 3. Method 4, syringe injection, was used with a low-cost manual HPCE instrument that is no longer manufactured (12).

The volume of material injected per unit time (V , nL/s) using methods 1-3 is determined by the Poiseuille equation (13)

V ^ = ^ ^ , (8.4) USrjL

where AP equals the pressure drop, D is the capillary internal diameter, 7] is the viscosity, and L is the length of the capillary. For gravity-based injections,

AP = pgAh , (8.3)

where p is the density of the sample solution, g is the gravitational constant, and Ah is the height difference between the liquid levels in the sample vial and in the detector-side buffer reservoir. Since the flow rate is proportional


to the forth power of the capillary diameter, these values can only be considered approximate. The specification for the variation of a 50-|Lim-i.d. capillary is ±1 |im.2 Table 8.3 contains data calculated for methods 1-3 for various values of the capillary i.d.

The consequences of an open-ended injection system and the Poiseuille equation mean that changes in the experimental conditions will result in variations of the amount of material injected. If the temperature is increased, more material enters the capillary because of the decreased viscosity of the BGE. If the capillary is lengthened, the amount of material injected decreases due to the increase in backpressure. This is important when contemplating methods transfer between instruments of different manufacture. Since the outlet side of the capillary often differs between brands, the injection time must be altered to compensate for this.

While exact assessment of the injected amount is generally unnecessary, since standards are used to calibrate the system, it is not difficult to calibrate the injection system. Such data can be useful in method validation, method transfer, and troubleshooting. The steps are as follows:

1. Fill the capillary with BGE and equilibrate to operating temperature. 2. Prepare a solution of BGE plus a UV-absorbing solute. Almost anything

that is soluble in the BGE and does not change viscosity can be used. 3. Set the injector pressure or vacuum to that specified in the method. 4. Inject the solution prepared in step 2 continuously until a baseline

deflection is observed.

^Polymicro Technologies.

Table 8.3 Injection Volume per Second for Pressure (0.5 psi), Vacuum (5" Hg), and Gravity (10 cm) versus Capillary i.d.

Injection Size (nL/sec)

Capillary Diameter (pm) Pressure

0.000078

0.039

0.60

3.0

9.6

Vacuum

0.00038

0.19

2.9

15

47

Gravity

0.000022

0.011

0.17

0.87

2.7

10

25

50

75

100

Capillary length: 100 cm; temperature: 30°C; viscosity: 0.801 gm cm'" s~ ; pressure injection: 0.5 psi (Beckman); vacuum injection: 5" Hg (Applied Biosystems); gravity injection, Ah = 10 cm (Waters) where g = 980 g cm-^ and p = 0.997 g mL-^


5. The flow rate of the injector is calculated as follows:

L(mm) ,^ ^. tiowrate(mm/s) = . Co.6)

t(s) Hydrodynamic injection is generally useful for capillaries with i.d.'s ranging

from 25 to 100 |Lim. For smaller i.d. capillaries, high-pressure injection must be used to keep the injection time reasonably short. For large-diameter capillaries, the injection pressure must be reduced to maintain an injection time of over 1 s. Shorter injection times may adversely affect precision.

The impact of the injection zone length on the electropherogram is illustrated in Figure 8.2. Since the sample was dissolved in BGE, the electropherogram with the 0.6-cm injection zone width (3% of the capillary) yields peaks that are much broader than desired. With the larger injections, the deterioration in efficiency is obvious. Using stacking electrolytes, 5-30% or more of the capillary can be filled with the sample.

Selection of pressure-driven versus vacuum-driven systems is not particularly important, unless interface to the mass spectrometer is required or high-viscosity polymer networks are employed. In these cases, the pressure-driven system is preferred. An advantage of vacuum-driven systems is that a perfect seal is required only on the outlet side. This holds for instruments that only employ an autosampler on the inlet. With pressure-driven systems, a perfect seal must be maintained over each sample vial, or injection errors will occur (14). This can be particularly important if vial closures are reused. Some instruments measure the pressure on the vial and adjust the injection time to compensate for any pressure variation.

B. ELECTROKINETIC INJECTION

The second injection technique uses electrophoretic and/or electroosmotic migration to inject samples into the capillary (13). By applying a low voltage for a short period, controlled amounts of the sample are introduced simply. As in the case with hydrodynamic injection, large (long) injections result in a substantial loss of resolution, particularly if the injection and run buffers are identical.

The quantity (Q) of a solute injected is given by

Q = (i ep + lijTtr^^Ct, (8.7)

where |iep and ii o are the electrophoretic and electroosmotic mobilities, respectively, r is the capillary radius, E is the field strength, t is the time of injection, and C is the concentration of each solute. This form of the equation is preferable to the simple volumetric relationship of Eq. (8.4), since solute discrimination may take place with electrokinetic injection (13, 15).


STABT

STOP

4.47

START

C 2.09

2.40

6.63

STOP

START

C 2.08

3 4.42

STOP

6.97

FIGURE 8.2 Impact of the injection zone width on electrophoretic efficiency. Capillary: poly-acrylamide-coated 20 cm x 25 im i.d.; buffer: pH 2.5 phosphate; solutes; substance P fragments dissolved in run buffer, 0.5 M-g/mL each; detection: UV, 200 nm. Injection width: top, 0.6 cm; middle, 2.0 cm; bottom, 3.0 cm. Reprinted with permission from J. Chromatogr., 480, 311 (1989), copyright © 1989 Elsevier Science Publishers.


High-mobility solutes may be preferentially enriched over those with low mobility This is illustrated in Figure 8.3. Solutes that have identical mobility in free solution show no such bias—for example, oligonucleotides and DNA fragments (16). This is fortunate, since it is often necessary to use electroki-netic injection with gel-filled capillaries or when high-viscosity polymer networks are employed.

Mobility bias is not a difficult problem to deal with. It can be compensated for by standardization or calculation, lonic-strength-mediated bias is a much larger problem. In Figure 8.4, a diagram of a capillary and electrode immersed in a sample is shown. The field strength at the point of injection is governed by the ratio of the conductivity of the BGE to that of the sample. If the sample has a high conductivity, two things occur. First, if the conductivity is due to the sample matrix, it becomes likely that matrix ions will enter the capillary in lieu of the solute. Second, the electric field is lowered at the point of injection, resulting in yet fewer ions entering the capillary. This type of bias was already

|a| Hfdrcittttlc lnjtcioii

10 m, 1 sec

250 260 440 450

Tifne |s)

600 610

FIGURE 8.3 Hydrostatic versus electrokinetic injection. Buffer: 20 mM MES adjusted with histi-dine to pH 6.0; solutes: Rb+, Li+, and arginine, 5 X 10"^ M; injection: (a) hydrostatic, Ah = 10 cm, t = 10 s; (b) electrokinetic, 1 s at 10 kV; detection: conductivity. Reprinted with permission from Anal Chem., 60, 375 (1988), copyright © 1988 Am. Chem. Soc.


ELECTRODE

FIGURE 8.4 Illustration of the electric field between the electrode and the capillary using elec-trokinetic injection. The field strength is influenced by the conductivity of the sample.

described in Section 6.1 ID. It is critical to maintain a constant and low conductivity in samples when using electrokinetic injection.

The problem with the electric field being influenced by the sample conductivity also affects the calibration curve. With the solutes dissolved in water, a negative deviation from linearity will occur at the higher solute concentrations. The calibration curves from hydrodynamic injection are linear up to the point where electrodispersion (Section 2.14) becomes significant.

While difficult in practice to employ quantitatively, electrokinetic injection can provide impressive trace enrichment. If the EOF is suppressed and the solutes are dissolved in water, it is possible to inject only ions without the bulk solutions. Those ions will stack at the head of the capillary, since the electric field strength is low. This topic will be covered in Section 8.6.

8.4 SHORT-END INJECTION

In previous sections, advantages of using the short end of the capillary for scouting runs were described. Another advantage of the short-end injection is enhanced sensitivity. Since the run time is short, diffusion-mediated band broadening is minimized (17, 18). To perform short-end injection, implement the following:

1. Equilibrate the capillary in BGE as usual. 2. Bring the sample to the capillary outlet. 3. Inject by pressuring the outlet vial or by electrokinetic injection using

negative polarity (inlet-side negative). 4. Set the power supply to negative polarity and perform the usual volt

age ramp.

8.5 Injection Artifacts: Problems and Solutions 331

On some instruments, it is necessary to use electrokinetic injection for short-end injections, because the instrument cannot provide low pressure on the outlet side.

Since the capillary length is short, the injection should also be kept small, and stacking buffers should be used. Be sure to set the detector time constant to 10-20% of the peak width to minimize that form of band broadening. Depending on the instrument, the short end of the capillary usually ranges from 6 to 10 cm.

8.5 INJECTION ARTIFACTS: PROBLEMS AND SOLUTIONS

In this section, problems unique to injection will be covered. Also included are the effects of damage to the inlet end of the capillary. A complete troubleshooting guide will be given in Section 10.8.

A. No INJECTION

The most frequent cause of no injection is a plugged capillary. This can result from evaporation of water at the capillary tip, which allows salt crystals to form. If the polyimide is not removed from the capillary tip, a shard of that material can enter and plug up the capillary. It is also possible that material from an unfil-tered sample or material that is insoluble in BGE can plug the capillary If the capillary is plugged, the observed current is usually zero or thereabout. Cutting a few millimeters of capillary from the end or replacing the capillary should remedy the situation.

No injection can also occur if an empty or incorrect sample vial is used, if an incorrect vial is called for in the method, if the vial cap is missing or badly leaking, or if the external pressure source is not activated. It is possible that the capillary is broken. Breaks usually occur at the detection window. Check that the voltage polarity is correctly set.

B. BAND BROADENING

This topic was covered in Section 8.1 as related to injection size. Working with the smallest practical injection size consistent with the required sensitivity is usually best, since sample matrix effects, particularly those related to high ionic strength, are minimized. If larger injections are required for sensitivity, it is best to inject low-ionic-strength samples to avoid "antistacking." Various stacking mechanisms will be covered in Section 8.6.


A slow injection sequence can cause diffusional band broadening (19), and so once injection is complete, the voltage should be immediately ramped up. Siphoning resulting from an imbalance of the fluid levels of the inlet and outlet reservoirs causes broadening as well. This is most critical for short injection zones, short capillaries, and wide-i.d. capillaries (19).

C. SPONTANEOUS (UBIQUITOUS) INJECTION

The simple process of inserting and withdrawing a capillary into a solution can cause an extraneous bolus of sample to enter the capillary. This effect was first noted in 1989 with the observation of a 3-nL (700-|Lim) injection on a 75-|im-i.d. capillary (20).

Large-diameter capillaries show greater spontaneous injection volumes. For example, a 75-|im capillary immersed in a dye solution for 20 s injected 4 nL of material. A 50-|im-i.d. capillary injected 2 nL of sample, which corresponds to a zone width of 1 mm. This is sufficient to yield a nonzero y intercept when a calibration curve is performed. Using a 25-|Lim-i.d. capillary, 0.1 nL was injected.

The mechanism for this effect appears to arise from an interfacial pressure difference formed at the inlet of the capillary when it is immersed in sample (21). Since the phenomenon is time dependent, it is critical to ensure that injection is performed in a repeatable manner, or else reproducibility will suffer. The effect of spontaneous injection is greatest when the size of the actual injection is small.

It has been shown that thin-walled capillaries can minimize this effect (21), but for the most part, spontaneous injection is ignored. There was one report of using spontaneous injection to perform small-volume injections, but the reproducibihty was 6% RSD (22).

D. SPONTANEOUS PEAKS

Spontaneous peaks can appear in either or both directions and always occur at the elution time of a neutral solute. The magnitude of the marker peak depends on the length of time the capillary is immersed in the electrolyte without applying the field. If the field is kept on, no additional spontaneous peaks are observed. These artifacts are due to undetermined effects of the physical geometry at the end of the capillary, which cause a concentration gradient to occur. These unusual peaks can sometimes be eliminated by trimming a small section of capillary or, failing that, replacement of the capillary. Since the peaks always occur at the migration time for the neutral marker, they can often be ignored. The use of buffers and reagents with low-UV absorbance will also serve to reduce this effect (23, 24).

8.5 Injection Artifacts: Problems and Solutions

E. PEAK TAILING

333

Though taihng can resuU from wall effects, there are also injection-related aspects. If the capillary is not cut squarely, a concentration gradient can occur upon injection (25). This is illustrated in Figure 8.5. This phenomemon highlights the importance of a properly prepared capillary.

0.03

AU

002

0,01

0.00

R

0,031-

AU

4 6 8

Migration Tima (mm) 10

0,02 h

0.01 i

000

1- B R

1 '

r S

\

^

1 „ , , 4 6 8

IVfigration Time (mm) 10

FIGURE 8.5 Effect of the physical shape of the capillary inlet on the resolution of naproxen enan-tiomers. (A) Properly cut capillary; (B) oblique cut (45°) capillary inlet. Reprinted with permission from And. Chem., 67, 2279 (1995), copyright © 1995 Am. Chem. Soc.


F. PEAK SPLITTING

Observation of split peaks is an unusual occurrence in HPCE, but there are numerous references describing the phenomenon (26-28). Here, some injection related causes of peak splitting will be described.

Splitting can occur when injecting solutes dissolved in organic solvents into MECC or cyclodextrin-containing electrolytes (29). This is probably due to the distribution of the solute between two phases moving at different speeds at the point of injection. The problem is resolved by dissolving the solute in aqueous media. If this is not possible, 6 M urea can be added both to the BGE and to the sample diluent to eliminate the problem.

Another, more subtle, cause of splitting is a fracture at the capillary tip. This can easily occur if the capillary hits a vial wall or seal. The polyimide coating keeps the cracked portion together. Upon injection, material moves into the capillary, both from the open end of the tube and through the crack. The split peak is usually smaller than the main component and always has a migration time that is a little shorter. If this occurs, feel the end of the capillary. If it is fractured, a small piece will usually come off, and the effect will disappear.

G. SAMPLE DEPLETION

When electrokinetic injection is employed, repeated injections from the same sample vial will show a continuing decline in peak height (25). This is particularly noticeable when injecting from very small sample volumes. This effect is partially due to the actual removal of solute ions from the sample, which decreases the solute concentration. The effect of buffer depletion is more important. This effect was described in Section 2.1.

With each injection, buffer ions electrophorese into the sample while the counterion migrates in the opposite direction. To maintain electrical neutrality, electrolysis also occurs, producing protons at the anode and hydroxide at the cathode. Since the sample serves as the electrolyte during injection, its pH and conductivity may change. As a result, it becomes more likely that ions other than the solute enter the capillary, and the driving electric field is reduced as well.

The problem is solved by injecting a water plug for a time equal to or slightly greater than the actual injection time. The water plug insulates the sample from the BGE. Buffer ions do not enter the sample if the plug is sufficiently large, and the water plug, an ion-depleted zone, serves to maintain a high electric field at the point of injection.

8.6 Stacking and Trace Enrichment 335

8.6 STACKING AND TRACE ENRICHMENT

A. INTRODUCTION

Compression, or stacking, of solutes in the injection zone is important for two reasons:

1. As Table 8.2 indicates, the injection size can severely limit the number of theoretical plates obtained for the separation.

2. The problems with detector sensitivity (see Chapter 9) can be solved in part by adequate trace enrichment of dilute solutes.

The problem posed by the first point is illustrated in Figure 8.6 using an MECC separation of urinary porphyrins (30). The injection size is only 10 nL into a 50 cm (length to detector) x 50 |lm i.d. capillary. In the electrophero-gram at left, the injection was performed with the sample dissolved in BGE, which contained 100 mM SDS and 20 mM CAPS, pH 11.

The sample was next dissolved in an electrolyte containing only 20 mM CAPS. Using the same BGE as before, the right-hand electropherogram was generated. Substantial improvements in resolution were found with the nonmicel-lar injection buffer. Removing the surfactant from the injection buffer reduced the ionic strength and, thus, the conductivity of the solution.

The second point can be addressed as well with a stacking electrolyte. Figure 8.7 shows the separations of some porphyrins with injection sizes ranging from 5 to 100 nL (30). A 100-nL injection represents about 10% of the volume of the capillary. Compared with a 5-nL injection, the limit of detection is improved by a factor of 13. Even with a stacking buffer, some injection-mediated band broadening can occur. While a 20-fold LOD improvement was expected, the improvement was only a factor of 13. For a 50-nL injection, the improvement is 8-fold (10-fold expected). Clearly, the LCDs can be improved at the expense of elec-trophoretic resolution. With proper optimization (see below), it is possible that both resolution and sensitivity can be further improved.

Stacking is not new in electrophoresis or even in HPCE. In 1979, Mikkers et a\. described stacking in CZE (31). In that work, they referenced analogous techniques using disk electrophoresis as far back as 1964. Stacking gels are widely used for many biochemical separations. In a gel-free medium such as CZE, stacking electrolytes can be employed to accomplish solute enrichment. As described in the following subsections, there are many mechanisms of and approaches toward stacking.

^Peak 1, mesoporphyrin, is not very soluble in the absence of the surfactant accounting for its diminished peak height.


III in

% m cc m

y o (0 UJ cc o 3

AGKMOUS INJECTION m F F e i 6 ,

lACELLAR INJECTION BUFFER

6

9

Uf^n*/NUA/w w v # / W

TIMEdnin.) FIGURE 8.6 Impact of injection buffer on electrophoretic resolution. Injection buffer: (A) 20 mM CAPS, pH 11, 100 mM SDS; (B) 20 mM CAPS, pH 11. Capillary: 72 cm (50 cm to detector) X 50 |im i.d.; BGE: 20 mM CAPS, pH 11, with 10% methanol; sample: porphyrin test mix, 5 nmoL/mL; injection: vacuum, 1 s; voltage: 30 kV; detection: fluorescence; excitation: 400 nm; emission wavelengths > 595 nm. Key: (1) mesoporphyrin (dicarboxyl); (2) coproporphyrin (tetracarboxyl); (3) pentacarboxyl porphyrin; (4) hexacarboxylporphyrin positional isomers; (5) heptacarboxyl porphyrin; (6) uroporphyrin (octacarboxyl). Reprinted with permission from J. Chromatogr., 516, 271 (1991), copyright © 1991 Elsevier Science Publishers.

B. IONIC-STRENGTH-MEDIATED STACKING

Differences between the conductivity of the injection zone and the BGE have an impact on the field strength that is distributed over each zone. As described in Section 2.1, this is simply a consequence of Ohm's law. Since a solute's electrophoretic velocity is proportional to the field strength, differing velocities can

TIME <miiiJ

FIGURE 8.7 Impact of injection size on resolution and sensitivity. Injection buffer, 20 mM CAPS, pH 11. Approximate injection size: (A) 5 nL; (B) 10 nL; (C) 25 nL; (D) 50 nL; (E) 100 nL. Other conditions as per Figure 8.6. Reprinted with permission from J. Chromatogr., 516, 271 (1991), copyright © 1991 Elsevier Science Publishers.


be realized within each compartment. The basis of stacking is to provide a high field strength over the injection zone. This is accomplished readily by injection of low-conductivity solutions.

The phenomenological situation at low pH values is illustrated in Figure 8.8. If a low-conductivity injection solution is employed, the field strength must be higher over that zone than over the remainder of the capillary. When the positively charged solutes migrate out of the injection zone and encounter the BGE, the field strength abruptly drops, and thus, the solute's electrophoretic velocity slows. Meanwhile, the solutes at the middle to rear of the injection zone are still exposed to the high field strength and continue to move forward at "full" speed. As a result, the ions in the injection band continue to narrow, until all have migrated into the BGE. The negatively charged counterions stack up as well, but at the rear of the injection zone (not shown).

At high pH values, as shown in Figure 8.9, a more complex situation occurs because of the presence of the strong EOF In this case, the EOF drives all solutes toward the cathode. The negatively charged anions migrate toward the anode and cross the boundary between the injection solution and the BGE at the rear of the injection zone.

Despite these differences in the pH-mediated direction of electrophoresis, the net result is band compression, the degree of which is related by the ratio of

LOW FIELD HIGH FIELD

ii)®(^©® a\:^'

FIGURE 8.8 Illustration of stacking of cations in a low-pH buffer.

8.6 Stacking and Trace Enrichment

LOW FIELD

339

HIGH FIELD

ep +

FIGURE 8.9 Illustration of stacking of anions in a high-pH buffer.

conductivities of the injection and separation electrolytes. These differences can be quantitatively expressed as (32)^

El

PI

(8.8)

where Ei and Ej are the field strengths over the injection zone and balance of the capillary, respectively, pi and p2 are the respective resistivities in these regions, Ci and Cj are the respective buffer concentrations, and / i s the field enhancement factor. Since the current or ion flux that passes through the capillary must be constant through each zone, the steady-state concentration of solutes [SJ and [S2] is inversely proportional to the field strength and, thus, the electrophoretic velocity; hence.

[SJVi ^ [ S j v ^ (8.9)

' The symbols describing the parameters given in (32) have been simplified for clarity.

3 4 0 Chapter 8 Injection

SO that

i ^ = l ^ = r , (8.10) [SJ [Q]

Since [SJ must increase, this can only be accomphshed by compression of the ions in the injection zone. Thus,

X3 = ^ , (8 .11) 7

where x^ is the effective sample plug length after stacking and x^ is the initial length of the injection plug.

Based on Eq. (8.11), the sample should always be prepared in water. Unfortunately, this is not optimal because of the generation of electroosmotic pressures. The measured EOF is based on the average electroosmotic contribution of each zone adjusted by the zone length. Since fluids can be considered incompressible, a hydrodynamic component is introduced whenever localized zones contribute differently to the total EOF This effect generates band broadening by contributing hydrodynamically enhanced diffusion. Calculation and plotting of these effects leads to Figure 8.10, which shows optimal results for field enhancement factors ranging from 5 to 20 (32). When yis small, the peak variance is proportional to the injection time. At high y values, the laminar-flow-induced broadening becomes significant.

These theoretical calculations are supported by experimental data shown in Figure 8.11 (32). The optimal procedure for stacking is to prepare the sample in an injection buffer that is 10-fold more dilute than the BGE. Based on Figure 8.7, it is possible to trade off some plates for sensitivity. Performed properly, ionic-strength-mediated stacking can improve sensitivity by a factor of 10.

At low pH, the low-ionic-strength injection buffer may still linger within the capillary and cause problems when very large injections are made (33). Other effects can be observed when using large-volume stacking hydrodynamic injections. Because the field strength over the point of injection becomes enormous, labile samples may decompose due to the generated heat (34).

C. IONIC-STRENGTH-MEDIATED ANTISTACKING

The antithesis of stacking is antistacking. The antistacking mechanism is illustrated in Figure 8.12 for cations at low pH. When a sample with a high ionic strength relative to the BGE is injected, the electric field over the injection zone declines as defined by Ohm's law. When a positive ion electrophoreses into the BGE, it becomes exposed to the high field strength over the BGE. As a result.


too Field enhancement factor Y

FIGURE 8.10 Plot of peak variance versus the field enhancement factor, y for several gravity-based (15-cm) injection times using a 100 cm x 75 jim i.d. capillary. Reprinted with permission from Ana!. Oitm., 63, 2042 (1991), copyright © 1991 Am. Chem. Soc.

the cation accelerates away from those cations, still remaining in the injection zone. The result is substantial band broadening.

Figure 8.13 shows the antistacking of anions at high pH. The anion crosses the boundary between the injection plug and the BGE at the rear of the zone. Now exposed to the high field strength, the anion accelerates toward the positive electrode. Ions remaining in the injection plug migrate more slowly toward the anode and, as a result, are pushed more rapidly by the EOF toward the cathode. Though the dispersion occurs at the opposite end of the injection plug than for cations, the ultimate result is the same.

Because of the phenomenon of antistacking, only small injections can be made when the ionic strength of the sample is high relative to the BGE. This same problem occurs in the slab gel (35). In that case, it was suggested to allow a timeout step after sample application, prior to applying the voltage. The timeout allows the rapidly diffusing salt ions to disperse from the sample zone. In the gel, 30 min was required for the timeout, and no diffusion-related band broadening of DNA was observed. While not reported in HPCE, a timeout may be possible, though not for 30 min. There have been several proposed schemes

c

3 -

3.

c

1 ' " <

J

1

iljj

6 minute

2 I

SP

6 minute Injection with pure water

4 -

c 3

3 -€ CO

J? 2 -c

-p , o 1 -«0 X2

<

6 minute in

1 1

1 1 I 1 1 1 1 k 1 1 JLA—/u \ ^

6 minute injection with 100 mM buffer

J \ ^ TV Migration time (min.)

FIGURE 8.11 Comparison of electropherograms of three different buffer concentrations in the sample plug. Capillary: 100 cm x 75 |im i.d.; buffer: 100 mM MES, pH 6.13; injection: gravity (15 cm) for 6 min (about 400 nL, or 5% of the total capillary volume); injection buffer: as specified on figure; voltage: 30 kV Key: (1) PTH-Arg; (2) PTH-His; (SP) sample plug; (I) impurity Reprinted with permission from Anal. Chan., 63, 2042 (1991), copyright © 1991 Am. Chem. Soc.

HIGH FIELD LOW FELD

•ep

-ep -eo

^+; t-' 'V

FIGURE 8.12 Illustration of antistacking of cations in a low-pH buffer.

8.6 Stacking and Trace Enrichment

HIGH FIELD

343

LOW FIELD

FIGURE 8.13 Illustration of antistacking of anions in a high-pH buffer.

that permit online desalting (36-38), though none are widely used. Voltage gradients shown applicable in CIEF (39, 40) have not been studied in CZE or MECC. In any event, it is often necessary to perform offline desalting to obtain optimal results.

D . pH-MEDiATED S T A C K I N G

For separations of zwitterions, the pH of the injection buffer can be advantageously selected to enhance peak compression (41). The stacking mechanism is shown in Figure 8.14. Using 10 mM ammonium hydroxide, peptides are injected into the capillary, which contains a 10 mM citrate, pH 2.5, supporting electrolyte. Upon application of the voltage, the negatively charged peptides migrate toward the anode. As the peptides at the rear of the band enter the acidic buffer, the charge flips and the direction of migration reverses. Meanwhile, the peptides at the front of the band are still migrating toward the anode. The result is the band collapses on itself. Finally, all the material in the narrowed band becomes positively charged and migrates toward the cathode. The small amount of injected ammonium hydroxide is dissipated and buffered.

The impressive stacking ability of this technique is shown in Figure 8.15 (41). The injection size was approximately 150 nL into a 72 cm X 50 |Llm i.d. capillary. Without the pH shift, no separation was obtained. With the pH shift.


LOW pH HIGH pH

IffjTcftSP' '""''''''"' '* "'' "'"*"' ''' ^

'p.^^?^:^i-'7S?^'^^?^M

CONCENTRATION • • • 0 -

SEPARATION

FIGURE 8.14 Illustration of pH-mediated stacking of zwitterions. Redrawn with permission from J. Chromaiogr., 516, 79 (1990), copyright © 1990 Elsevier Science Publishers.

the resolution is comparable to a 5-nL injection (data not shown). The limit of detection for some dynorphins can be less than 10 ng/mL using a 300-nL injection (data not shown). This technique has been applied toward screening of narrow-bore LC fractions of peptide maps. No data has been published as of this writing on the repeatability of this technique.

A variant of pH-mediated stacking can be used for DNA separations including sequencing (42). In this case, electrokinetic injection of hydroxide can be used to neutralize the Tris buffer commonly used for DNA separations. This creates a low-conductivity zone where the DNA fragments are enriched.

E. ACETONITRILE-SALT MEDIATED STACKING

A stacking technique with an unknown mechanism occurs when mixtures of acetonitrile and high concentrations of salt are present in the sample (43-46). The technique is useful when analyzing drugs in blood serum or plasma, since the acetonitrile can be used to deproteinize the sample and simultaneously provide the stacking environment. Urine can be employed as a sample as well. The best results are found when dissolving the sample in two parts acetonitrile and one part sodium chloride (43). As much as 50% of the capillary can be filled with the sample. A 250 mM boric acid buffer, pH 8-9, appears to give better results than phosphate buffer, presumably due to the lower conductivity of boric acid. Stacking is observed in the absence of SDS or cyclodextrins, unlike the situation described in the next subsection.


^

-' H^ TiME(min.}

FIGURE 8.15 Separation of peptides using pH-mediated stacking. Capillary: 72 cm (50 cm to detector) x 50 |Lim i.d.; separation buffer: 10 mM citrate, pH 2.5; injection buffer: 10 mM ammonium hydroxide; injection: vacuum, 30 s (about 150 nL); detection: UV, 215 nm; temperature: 30°C; solutes: dynorphins, 10 |ig/mL. Top: injected in separation buffer; bottom: injected in injection buffer. Redrawn with permission from J. Chromatogr., 516, 79 (1990), copyright © 1990 Elsevier Science Publishers.

F. STACKING OF NEUTRAL SOLUTES

Several schemes have been reported for the stacking of neutral solutes during MECC separations (47-52). Some of these stacking mechanisms are covered in this subsection.

Since neutral solutes do not migrate via electrophoresis, charged reagents must be used to facilitate stacking. If the solute is dissolved in a diluted version of the BGE, then the field strength over the point of injection is high. The charged micelle within the injection zone then migrates quickly, until it encounters the BGE. Solutes attached to the injection micelle are thus enriched. In this form of stacking, solutes that spend more time attached to the micelle show


greater enrichment factors than those spending more time in the bulk water (51). Other reports indicate enrichment is independent of retention (50) when the solute is dissolved in water. Like stacking of charged species, this mode provides an order of magnitude of trace enrichment and is simple to implement.

More sophisticated schemes can yield yet higher enrichment factors. At pH 2.5, the net migration velocity of an SDS micelle is greater than the EOF, and so the surfactant migrates toward the positive electrode; thus, reversed polarity is employed (Section 4.4). Injection can be hydrodynamic (49), or if a water plug is preinjected, electrokinetic (53).

The situation is diagrammed in Figure 8.16. Upon application of the field, the negatively charged micelle migrates into the injection zone, where it becomes exposed to the high field. The micelles stack and thus enrich the neutral solute. As the surfactant concentration increases, the field strength declines until it equals that over the BGE, stacking ceases, and the separation proceeds. In the electrokinetic injection mode, hydrophobic solutes such as steroids can be enriched by a factor of 50, whereas hydrophilic solutes such as phenols are enriched by a factor of 3 (53). The hydrodynamic injection mode appears more effective for hydrophilic species; phenol was enhanced by a factor of 13, and 2,3,5-trimethylphenol was concentrated 131-fold (49).

LOW FIELD HIGH FIELD

WATER PLUG • OR • DILUTE SAMPLE G-^

FIGURE 8.16 Stacking of neutral molecules by MECC. • represents a neutral molecule, whereas the negative ions represent anionic micelles.

Very large injections can be made using these techniques. Hydrodynamic injection of 10,000 mbs has been reported. For electrokinetic injection, a water plug up to 8.8 cm is preinjected, followed by electrokinetic injection until the current reaches 70% of that of the BGE by itself. Overinjection is indicated by the onset of peak splitting.

A novel twist for the stacking of neutral molecules employs a high-ionic-strength injection solution (54). Figure 8.17 illustrates the stacking of some

o as

<

Migration Time (mia)

FIGURE 8.17 Influence of salt concentration on the stacking of steroids. BGE: 80 mM cholate, 10 mM borate, 10% ethanol; capillary: 47 cm (40 cm effective length) x 50 jim id; injection zone: 3.6 cm; voltage: 30 kV; detection: UV, 254 nm. Key: (1) cortisone; (2) Cortisol; (3) 11-deoxycorti-sol; (4) 17a-hydroxy-progesterone; (5) progesterone. Reprinted with permission from Anal. Chem., 1999; 71:1679, copyright © 1999 Am. Chem. Soc.


neutral steroids using MEKC with 80 mM sodium chelate, 10 mM borate, 10% ethanol as the BGE. As the salt concentration in the sample is increased beyond 50 mM, the peaks sharpen dramatically. It turns out that 50 mM sodium chloride is equal in conductivity to the BGE.

The mechanism for this stacking effect is shown in Figure 8.18. Because of the high salt content in the sample, the electric field over the injection plug is reduced relative to that over the BGE (Figure 8.18, top). At high pH, the EOF is directed toward the cathode and pushes everything in that direction. The countermigrating negatively charged surfactant micelles enter the front of the injection plug and become exposed to the low field strength in that region. As a result, their countermigration toward the anode slows, and they stack up at the head of the zone. The neutral steroids are pushed toward the cathode by the FOE At the front of the injection zone, they encounter a very high concentration of cholate micelles (Figure 8.18, middle). Since the steroids bind to the micelle through hydrophobic interaction, the solute concentration builds up at the zone boundary as well.

Not shown in Figure 8.18 are the sodium and chloride ions. The mobihties of these ions are very high, and as a result, they migrate rapidly from the injection plug and soon end up in their respective electrolyte reservoirs. Once the salt ions depart the injection plug, stacking is complete, and the now-enriched steroid zone (Figure 8.18, bottom) continues down the capillary as a result of the EOF

HIGH FIELD LOW FIELD

FIGURE 8.18. Diagram illustrating the mechanism of high-salt mediated stacking in MECC. represents a neutral molecule, whereas the negative ions represent anionic micelles.


The significance of this approach is the abihty to stack in the presence of sak. This solves in part an ongoing problem in HPCE: how to deal with samples having high salt concentrations. While reduced in practice to neutral solutes, the technique should work well with cations that can ion-pair to the surfactant. To stack anions, a cationic surfactant would be required. The technique works as well when charged cyclodextrins are present, and so the method can be extended to chiral separations in high-concentration salt environments.

G. WHOLE-CAPILLARY INJECTION

Very large injections can be performed by removing the injection solution from the capillary after stacking (33). For example, the entire capillary or a large portion thereof can be filled with an anionic sample dissolved in water (Figure 8.19, top). When a negative voltage is applied, the anions migrate toward the positive electrode and stack at the boundary between the injection zone and the entering BGE (Figure 8.19, middle). The EOF naturally drives the overall migration velocity toward the negative electrode. Simultaneously, the water elec-troosmotically exits the capillary at the negative-electrode injection end. As water exits, the BGE enters the capillary at the opposite end. As the analyte encounters the BGE, migration ceases; this cessation occurs because there is virtually no field strength over the high-ionic-strength BGE, since the voltage drop occurs over the water. When the water is nearly out of the capillary as shown by the current reading, the power supply polarity is reversed, and the analytes are quickly separated and eluted (Figure 8.19, bottom).

^ . ._ 3eeeee eeeeeeett'eeeseeeeeeee

• vwewee Sx-)60B66

* — E O F EP-

FIGURE 8.19 Illustration of the whole-capillary injection process for anions.

This process is illustrated in Figure 8.20 using some PTH-amino acids prepared in water at the CLOD for a normal run (33). Since the entire capillary is filled with sample, the baseline is elevated slightly as the run begins, indicating the presence of the analyte. Stacking occurs at the rear of the zone, and a peak is seen as the stacked but unseparated anions pass the detector for the first time, driven by the EOF. When the current rises to 99% of the value for the BGE (previously determined), the electrode polarity is reversed. The separation occurs on the segment of capillary that was beyond the detector window prior to the switching of the power supply polarity

A similar process can be performed for cations, provided a surfactant is added to reverse the EOF (33). While there are no data on the precision of this technique, the process may be useful for micropreparative separations; however, ionic strength variations between samples will badly degrade this mode of stacking. This technique has not been used widely, perhaps for this reason.

4.0 H

^ 3.6-1 c 3 jQ 3.0

0) 2.5-] C m -£ 2.0 o (A

JDi < 1.6-^

1.0

negative species

water plug

, switched electrodes

I—B

lil ,j^ . J p. 6 10 15

Migration Time (min) 20

FIGURE 8.20 Electropherogram from a whole-column injection. The whole capillary [100 cm (65 cm to detector) x 50 |im i.d.] is filled with sample, (A) PTH-Arg, 4.6 x lO'^ M; (B) PTH-Glu, 3.4 X 10"^ M dissolved in water. The ends of the capillary are dipped in support buffer, 100 mM MES adjusted to pH 6.1 with 100 mM histidine. At 4.5 min, all anions passed the detector as a single band. At 7 min, the polarity is reversed, and the anions migrate back in the direction of the injection end and pass the detector as separated analytes. Reprinted with permission from Anal. Chem., 64, 1046 (1992), copyright © 1992 Am. Chem. Soc.


H. STACKING WITH ELECTROKINETIC INJECTION

Chien and Burgi (55-58) introduced the term/ield-ampli/ied injection (FAI) for describing stacking with an electrokinetic injection. This is quite descriptive, since the electric field is indeed amplified over the injection zone when low-conductivity injection solutions (as compared with the supporting electrolyte) are employed.

To perform FAI, the sample must be dilute and dissolved in water. The capillary is filled with the supporting electrolyte, and a small plug of water (10 nL) is hydrodynamically injected at the head of the capillary. This water plug is to ensure a high electric field at the point of injection. Sampling bias characteristic of electrokinetic injection is reduced, though not eliminated, with the water plug as shown in Figure 8.21 (55). Following introduction of the water plug, the capillary-electrode assembly is immersed in the sample, and a voltage is applied for injection. Injection voltages as high as 30 kV can be employed.

The mechanism of peak compression is similar to that in ionic-strength-mediated stacking. As the front of the zone enriches, the local ionic strength increases, lowering the field strength in that region. Solute at the rear of the band is still exposed to a high field strength and continues to migrate at full speed, until it encounters the higher ionic strength region. The water plug differentiates this form of peak compression from conventional stacking; the stacked solute provides the higher ionic strength region.

The length of the injection plug using FAI can be estimated from

X, = ( ^ + / i , j E t , (8.12) r

where r is the ratio of the resistivity of the supporting electrolyte to that of the injecting solution. For conventional electrokinetic injection, the expression simplifies, since r = 1. Thus, in FAI, the impact of the EOF on the band width is reduced, and so higher voltages or longer injection times can be employed without excessive band broadening.

Representative electropherograms are given in Figure 8.21 for (top) hydro-dynamic injection, (middle) electrokinetic injection with the sample prepared in water, and (bottom) electrokinetic injection using a water plug in front of the aqueous sample. An LOD of 10~^ M for PTH-Arg has been reported (55).

A consequence of FAI is selective ion injection (56). It is possible to enhance selectivity and speed separations by introduction of either cationic or anionic species exclusively. For positive-ion electroinjection, the sample side of the capillary is held at positive polarity. Nearly all of the electric field is placed over the water and sample zones; thus, only cations electrophorese into the capillary, and the anions migrate toward the positive electrode and never enter the capillary.

T '^ a

>>.^;JU^»V^'*^*^'*^" ^h^^^^^^^^^S^'^f'i^h^

5 10

Time Cmin)

A

< o

—i 1 . . I l _ —i i i—

5 10

Time (min)

C

5 10 1

Time Cmin)

FIGURE 8.21 Separation of PTH-amino acids with injection by (top) conventional electroki-netic injection with sample dissolved in the running electrolyte; (middle) electrokinetic injection with sample dissolved in water; (bottom) field-amplified injection where a small plug of water is introduced into the capillary prior to electrokinetic injection. All electrokinetic injections were at 5 kV for 10 s. Capillary: 100 cm (75 cm to detector) x 75 |J.m i.d.; buffer: 100 mM MES-HIS, pH 6.2; voltage: 30 kV. Solutes: (A) PTH-Arg; (B) PTH-His; (C) neutral marker. Reprinted with permission from J. Chromatogr., 559, 141 (1991), copyright © 1991 Elsevier Science Publishers.


Since the field strength is carried over the narrow injection zone, the EOF that is averaged over the entire capillary is very small.

To inject negative ions, a water plug is injected, followed by the sample using negative polarity. After injection, the capillary is returned to run buffer, and the polarity is reversed to sample-side positive. The anions quickly encounter the run buffer as they electrophorese toward the positive electrode. The field strength over the sample zone decreases, so that the anions do not reenter the electrode reservoir. As the water plug mixes with buffer, the field strength redistributes evenly over the entire capillary, and the separation proceeds as usual.

To inject both cations and anions, inject the water plug, and then, with positive polarity, electroinject the cations. Switch the polarity to negative to elec-troinject the anions, return the capillary to the run buffer, and reverse the polarity again to run the separation. For all these injection methods, FAl stacking is in full force because of the difference between the ionic strength of the running electrolyte and that of the injection solution.

The merits of these FAI techniques are illustrated in Table 8.4 and compared with those of gravity injection and conventional electrokinetic injection. Substantial discrimination is obtained using either the positive- or negative-ion mode. Using the polarity-switching technique for introduction of both cations and anions, the technique is biased toward cations. Like most of these stacking modes of injection, the precision has not been adequately studied. The sample matrix can have an extreme effect on both the precision and the quantitative accuracy of these techniques. It is extremely important that a consistent injection matrix be employed.

This technique has been improved on over the years (59-61). Rather than diluting the sample with water, 80% propanol containing 100 |LlM phosphoric acid better controls the distribution of the electric field. The preinjection water zone is 1 mm. Injections of no longer than 90 s at 20 kV should be used to prevent capillary damage. After performing solvent extraction, evaporation, and

Table 8.4 Peak Heights^ for Anions and Cations Using FAI Compared with Conventional Injection Techniques

Gravity injection

Electroinjection'^

FAI—positive-ion mode

FAI—negative-ion mode

FAI—dual-ion mode

PTH-ARG

1

0.311

28.04

0

32.03

PTH-HIS

1

0.225

13.44

0

9.28

PTH-ASP

1

0.025

0

13.66

2.58

PTH-GLU

1

0.022

0

12.58

1.96

^Peak heights normalized to gravity injection. ''Injected using the run buffer, 100 mM Mes-His. Data from J. Chromatogr., 1991; 559:153a.


reconstitution in 200-|lL propanol-phosphoric acid solution, it is possible to electroinject all of the ions in the sample and reach sensitivities comparable to those of HPLC. Further improvements can be made by injecting a 2-6 mm plug containing 200 mM sodium phosphate in 90% ethylene glycol prior to the water plug. This solution serves to stack or slow the ions after they leave the pure water plug. The ion-trap solution has high viscosity and conductivity (relative to the BGE), both of which reduce electrophoretic mobility With the use of an internal standard, superior precision compared to HPLC is often obtained.

I. TRANSIENT ISOTACHOPHORESIS

Commercial instrumentation for capillary isotachophoresis (CITP), also known as displacement electrophoresis, has been available since 1974 with the introduction of the LKB Tachophor, the Shimadzu IP-IB, and later, the Spioska Nova Ves (Czechoslovakia) CS ZKI 001. Despite a wealth of commercial instrumentation and extensive theory and applications, CITP did not become a routine separation tool in most analytical laboratories. This is particularly true in the United States. In parts of Europe, CITP has become an important technique, especially in the Czech Republic.

Though CITP is used much less than CZE, a stacking technique known as transient ITP or tITP has important applications. This technique permits the concentration of trace components while leaving the major component unen-riched. Trace enrichment of up to lOOOx is possible. In order to understand tITP, the general principles of ITP must first be considered.

Isotachophoresis literally translates to "electrophoresis at uniform speeds." This means that the transit time of a solute through the capillary under iso-tachophoretic conditions is independent of mobility To understand this concept and its practical implications, some arguments described in Section 2.1 will be employed.

The separation scheme of CITP is illustrated in Figure 8.22. Unlike CZE, the buffer system for CITP is heterogeneous. Before beginning a run, both the inlet and outlet buffer reservoirs are filled with a leading electrolyte, or leader. The leader is selected to have a mobility greater than that of any of the components to be separated. The capillary is then filled with the leader.

For example, when separating anions by anionic CITP, the highly mobile chloride ion is most frequently selected as the leader. A large-volume injection (up to 50% of the capillary) is then performed. The inlet reservoir is changed to a terminating electrolyte (or terminator), the mobility of which is less than that of any of the components in the sample mixture. The voltage is then applied.

The electrophoretic velocity is given in

Vep = MepE- (8.13)


v i i i l l i i i i i i ^ ELECTROLYTE

SAMPLE l i l i i i i l i i i i i i

DIRECTION OF SEPARATION

FIGURE 8.22 Schematic representation of a three-component separation by CITP at the moment of injection (top) and after separation (bottom).

Since the buffer system and the sample are heterogeneous, the field strength will be inversely proportional to the conductivity of each individual zone. The conductivity depends on the mobility and the concentration of each solute. Unlike CZE, CITP bands are always in contact with each adjacent zone. This feature is necessary to maintain electrical continuity throughout the system, since there is no supporting electrolyte.

As a first approximation, assume the concentrations of the leader, terminator, and sample components are identical. As the run proceeds, individual solutes begin to migrate as determined by their individual mobilities. As the bands begin to sort out, the field strength over each sample zone begins to change. Highly mobile sample components are also highly conductive. As a result, E and thus Vgp are reduced for highly mobile bands. Conversely, sample components with low mobility generate greater field strengths, thereby increas-ii g ep- When the system reaches equilibrium, each solute's electrophoretic velocity becomes identical. The isotachophoretic velocity at equilibrium for a single-component separation can be expressed as

ViTP = ELML = EsMs = ETMT , (8.14)

where |X = the respective mobilities of leader, solute, and terminator, and E = the field strength over each isotachophoretic zone. At the steady state, all zones move at the same speed and exhibit stable, well-defined boundaries. This feature is illustrated in the bottom section of Figure 8.22 for a three-component system.

Let us add some complexity to the illustration with a two-component sample. Solute 1 has high mobility but low concentration. Solute 2 has low mobility


and high concentration. This is illustrated in Figure 8.23A. Upon activation of the voltage, solute 1 begins to migrate ahead of solute 2. By definition, solute 1 cannot enter the leader zone by electrophoresis, since it is of lower mobility If some should enter that zone by diffusion, the solute is exposed to a low electric field and thus slows down. As solute 1 enriches toward the front of the zone, the field strength rises, as the solute is dilute, and stacking occurs. On the other hand, solute 2 cannot stack, since it is already quite concentrated and generates a low electric field strength. These concepts are illustrated in Figure 8.23B. Now the importance of CITP becomes apparent. Only dilute components are enriched. In conventional ionic-strength-mediated stacking, all components are enriched.

The concentration of all CITP bands is related to the concentration of the leading electrolyte as described by the Kohlrausch regulating function

U L ^ A + Mc)J Q = Q (8.15)

where Q = analyte concentration at steady state; Q = concentration of the leader; and the mobility terms are for the analyte, the leader, and the leader's counterion. Since the concentration of the leader (usually 10 mM) is selected to be much higher than the solute concentrations, band compression always occurs because of the high field strength over the solute zones.

Since the peak capacity of CZE exceeds that of ITP, the goal was to couple the techniques together to realize the best features of both. Early work in this field employed two capillaries that were coupled together, one for ITP and one for CZE (62, 63). Since special instrumentation was required, this technique was quickly abandoned, and single-capillary ITP-CZE was developed.

B DIRECTION OF SEPARATION

L +

FIGURE 8.23 Diagram of a two-component CITP separation where/is > Ms and [S2] > [SJ.


The goal is to employ ITP only briefly during the injection process for enrichment, followed by separation via CZE. In doing so, CITP is employed only for a short time, and then the conditions for CZE are restored. In other words, a heterogeneous buffer (leader-terminator) system exists transiently, followed by restoration to the homogeneous environment of CZE. There are many approaches for meeting these requirements (64-75).

First, a good CZE separation must be developed. The only new requirement is that the buffer co-ion must serve as a leader or a terminator. A series of ITP electrolyte systems is given in Tables 8.5 and 8.6. If a leader is selected, then fill the capillary with that buffer, inject 10-50% of the capillary with sample, and immerse the capillary in a terminating buffer. Activate the voltage for 30-90 s, and then return the inlet side of the capillary to the leading electrolyte and continue the run. In this example, after the focusing step, the leading ions quickly overtake the terminator and sample ions, and now a leader-sample-leader situation is regenerated, as required for CZE.

In another mode, leader or terminator is added to the sample itself. If a sufficiently large injection is made, the conditions for ITP are fulfilled for a short time prior to the system automatically converting to CZE. If you are seeking to enrich trace components in the presence of a major component, the major component itself can serve as a leader or terminator. This is termed sample self-stacking (64, 65).

Isotachophoresis can occur by accident if any of the foregoing conditions are met (30, 76). This is illustrated in Figure 8.24 using the urinary porphyrins applications described in Section 4.8. The injection is a urine sample from an

^Provided by Vladislav Dolnik from the CZE-ITP Internet Discussion Group.

Table 8.5 Buffers for Anionic ITP^

pH Base Terminator

3.6

4.3

4.9

6.1

7.1

8.1

9.5

P -Alanine

EACA^

Creatinine

Histidine

Imidazole

Tris

Ethanolamine

Glutamic, nicotinic, or pivalic acid

Pivalic acid

IVIES'

IVIES'

Hepes' + barium hydroxide

Glycine + barium hydroxide

EACA^ + barium hydroxide

^Use 10 mM hydrochloric acid (chloride as leader) with 20 mM base. ^'e-Amino-N-caproic acid. *^2-(N-Morpholino)ethanesulfonic acid. '^N-Cl-Hydroxyethyl) piperazine-N'-(2-ethanesulfonic acid).


Table 8.6 Buffers for Cationic ITP^

pH

4.7

6.1

7.2

8.1

8.7

9.8

10.3

Base

Acetic acid

MES^

MOPS^

Tricine

Asparagine

Glycine

/^-Alanine

Terminator

Acetic acid (proton as terminator)

Histidine

Imidazole

Tris or triethanolamine

Tris

Ammonium hydroxide

Ammonium hydroxide or methylglucamine

^Use 10 mM potassium hydroxide (potassium as leader) with 20 mM acid. '^2-(N-Morpholino)ethanesulfonic acid. ''3-(N-Morpholino)propanesulfonic acid.

individual not having a porphyria. In Figure 8.24A, note the appearance of a spike for peak 6, uroporphyrin. This spike corresponds to milhons of theoretical plates. The spike was repeatable and thus not attributable to arcing or specks of material.

The sample was next fortified with porphyrins, and again, only the uroporphyrin is focused (Figure 8.24B). Upon reduction of the injection size (Figures 8.24C and D) the uroporphyrin peak broadens, whereas the other peak widths remain the same.

It is likely that endogenous chloride from the urine serves as the leading electrolyte and the mechanism of stacking is tITP Since only uroporphyrin is focused, the terminating ion (unknown, perhaps SDS or CAPS from the BGE) must have a mobility lower than that of uroporphyrin but greater than that of any of the other porphyrins. When a small injection is made, there is insufficient time for the chloride to set up as a transient leading zone prior to its migration toward the inlet (positive electrode) and out of the capillary While the solutes elute toward the negative electrode (by the EOF), electrophoresis of the anions is toward the anode, and this is anionic ITP Proof that tITP is occurring is obtained by increasing the injection size and observing a sharpening of the focused peak(s).

J. PRACTICAL ADVICE

Considering that tITP is an advanced procedure and not for the faint of heart, it is critical for this and all stacking techniques to carefully control the conductivity of the sample. With tITP, this is particularly important, since the injection size is so large. Regardless of which stacking technique is employed, smaller injections always provide for a more robust separation. When selecting a stacking procedure, it is advisable to start with the simpler methods, such as ionic-strength-mediated stacking.


MATRIX EFFECTS * UHtNi

D SPiKE, 2s

C SPIKE, 3s

W*WJ' w 'f»^

B SPIKE,

, ^ J ^ ^ ^ 0 i / ^ ^ ^ ^ MS^

16

TlfVIECmln.)

FIGURE 8.24 Impact of injection time on the tITP of urinary porphyrins. Conditions and key as per Figure 8.6. Sample: (A) urine from a porphyria negative individual; (B-D) urine spiked with 300 pmol/mL porphyrins. Injection times as specified on figure. Reprinted with permission from J. Chromatogr., 516, 271 (1990), copyright © 1990 Elsevier Science Publishers.

The size of the injection is dictated by the requirements of the Umit of detection. It is often prudent to employ extended path length capillaries and offline sample preparation to help meet the required LOD. This eases the requirements for stacking and provides for more stable separation conditions.

K. MEMBRANE AND LC-BASED ENRICHMENT

The development of a device for online trace enrichment has not been straightforward. A few years ago, a capillary containing a 1-2 mm plug of a polymeric


reversed-phase packing was commercially available, but it no longer is (Jl). Large volumes of aqueous injection buffer can be loaded into the capillary by electrokinetic injection. Enrichment occurs through binding of hydrophobic solutes to the polymeric packing. After loading, a small volume of organic solvent is injected to elute the solutes into the capillary, after which CZE is performed in the usual manner. CLODs for peptides as low as 1 ng/mL have been reported, with migration time and peak area precision of better than 1.5%.

There is ongoing research into the development of a membrane-based device to provide for online trace enrichment (78-84). These membranes allow injection sizes of 1 |LiL. Solutes are enriched by hydrophobic interaction with the membrane. Such a large sample injection would solve most sensitivity problems in HPCE, but these membranes are not commercially available. They remain a research tool.

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65. Gebauer, P., Thormann, W, Bocek, P. Sample Self-Stacking and Sample Stacking in Zone Electrophoresis with Major Sample Components of Like Charge: General Model and Scheme of Possible Modes. Electrophoresis, 1995; 16:2039.

66. Foret, E, Sustacek, V, Bocek, P On-Line Isotachophoretic Sample Preconcentration for Enhancement of Zone Detectability in Capillary Zone Electrophoresis. J. Microcolumn Sep., 1990; 2:229.

67. Foret, E, Szoko, E., Karger, B. L. On-Column Transient and Coupled Isotachophoretic Preconcentration of Protein Samples in Capillary Electrophoresis. J. Chromatogr, 1992; 608:3.

68. Foret, E, Szoko, E., Karger, B. L. Trace Analysis of Proteins by Capillary Zone Electrophoresis with On-Column Transient Isotachophoretic Preconcentration. Electrophoresis, 1993; 14:417.

69. van der Schans, M. J., Beckers, J. L., Moiling, M. C, Everaerts, E M. Intrinsic Isotachophoretic Preconcentration in Capillary Gel Electrophoresis of DNA Restriction Fragments. J. Chromatogr, A, 1995; 717:139.

70. Witte, D. T, Nagard, S., Larsson, M. Improved Sensitivity by Online Isotachophoretic Preconcentration in the Capillary Zone Electrophoretic Determination of Peptide-like Solutes. J. Chromatogr, A, 1994; 687:155.

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78.Benson, L. M., Tomlinson, A. J., Mayeno, A. N., Gleich, G. J., Wells, D., Naylor, S. Membrane Preconcentration-Capillary Electrophoresis-Mass Spectrometry (mPC-CE-MS) Analysis of 3-Phenylamino-l,2-propanediol (PAP) Metabolites. J. HighResolut. Chromatogr, 1996; 19:291.

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CHAPTER 9 Detection

9.1 On-Capillary Detection 9.2 The Detection Problem 9.3 Limits of Detection 9.4 Detection Techniques 9.5 Band Broadening 9.6 Absorption Detection 9.7 Fluorescence Detection 9.8 Derivatization 9.9 Mass Spectrometry 9.10 Micropreparative Fraction Collection

References

9.1 ON-CAPILLARY DETECTION

The most common mode of detection in HPCE is on-capillary detection. Considering the minuscule dimensions of the capillary, this mode of detection has two advantages:

1. Since the capillary is contiguous between the electrodes, current leakage is eliminated.

2. Dilution of the eluting solutes either from dead volume in a flowcell or from the postcapillary reagent or sheath flow is eliminated as well.

The characteristics of on-capillary detection differ dramatically from those of postcolumn detection in HPLC. In chromatography, solutes move through the chromatographic packing at velocities determined by the mobile-phase flow rate and the overall retention characteristics of each analyte. On the column, the peak velocities depend on each solute's retention characteristics. Once off the column, all solutes are swept past the detector by the mobile phase at identical flow rates. The detected peak widths are a function of chromatographic processes and are not related to the peak velocity past the flowcell.

365

366 Chapter 9 Detection

In HPCE, a different set of rules applies (1). The migration velocity of each solute through the capillary is a function of its electrophoretic mobility in conjunction with the EOE Since detection occurs on-capillary, these forces are operative as the solute is traversing the detection window. As a result, slower moving components spend more time migrating past the detector window than their more rapidly moving counterparts.

Figure 9.1, top trace, illustrates the separation of a three-component mixture recorded directly from the detector output (1). The bottom trace gives the same separation, corrected for the zonal velocity. This correction is calculated by

(9.1)

where Wg is the spatial width of the sample in units of length, L is the effective capillary length, t^ is the migration time, w is the recorded temporal width in time units, and w^ is the spatial width of the detector window. Thus, two elec-tropherograms can be defined. The spatial electrogram refers to the actual band-

- i i DETECTOR INJECTOR

SPATIAL i ELECTROPHEROGRAM |

TEMPORAL ELECTROPHEROGRAM

TIME OR LENGTH

FIGURE 9.1 Plots of the detector response (bottom trace) as a function of time and (top trace) as a function zone length within the capillary. Redrawn with permission from J. Chromatogr., 480, 95 (1989), copyright © 1989 Elsevier Science Publishers.

9.2 The Detection Problem 3 6 7

width on the capillary, whereas the temporal electrogram is defined by what the detector observes.

This phenomenon has practical implications, since slower moving zones produce an increase in the peak area counts. When quantifying solutes by a response factor, a correction factor must be applied to normalize the peak area irrespective of the migration velocity:

Aeo. = ^ ^ , (9.2)

Here, Aaw is the measured peak area, t^ is the migration time, and A orr is the corrected peak area.

Response factors are frequently employed when standards are unavailable. Applications involving oligonucleotides, impurity analyses of drugs, and chiral separations are typical examples. For these applications, it is assumed that all solutes have the same molar absorptivities.

Whenever quantitation without a matching standard is used, it is necessary to normalize the peak areas. While the peak height is not related to the solute velocity as it passes through the optical window, quantitation is less accurate than peak area because of stacking/antistacking and electrodispersive effects.

When standards are used, it is unnecessary to provide this correction, since it is assumed that the standard behaves identically to the solute, if the migration times are constant. If the migrations times vary slightly from run to run, area normalization does not improve precision (2). If the migration time precision is poor, it is best to correct that problem rather than normalizing the peak areas.

9.2 THE DETECTION PROBLEM

Because of the minute amounts of material injected into the capillary, extremely high sensitivity detection is generally required for all forms of HPCE. The problem is exacerbated by the desire to dilute samples to rid separations of troublesome matrix effects. The instrumental problems for optical detection are twofold: (i) the short optical path length as defined by the capillary i.d.; and (ii) the poor optical surface of the cylindrical capillary.

While square and rectangular capillaries have been around for some time, there have been no definitive studies showing any advantages in detection. An exception to this is viewing down the long end of a rectangular capillary (3). This approach resulted in substantial band broadening. In addition, proper alignment of square and rectangular capillaries in the optical window is critical.

Commercial instruments use absorption detectors that are modifications of standard HPLC detectors. The absorption detectors are modestly sensitive, giving limits of detection of 10"^ M for solutes with very high molar absorptivities. While laser fluorescence detection can improve sensitivity down to 10" ^ M with commercially available equipment and approach single-molecule detection in


more sophisticated apparatus, derivatization is usually required to tag a solute with an optimized fluorophore.

Innovation is still required to solve the general detection problem in HPCE. Improvements of two to three orders of magnitude in absorption detection would solve many problems relating to matrix effects and Unear dynamic range. With the high resolving power of HPCE, less-selective detectors might prove useful, providing the sensitivity requirements are fulfilled.

9.3 LIMITS OF DETECTION

There are two means of describing the limits of detection of a system: the concentration limit of detection (CLOD) and the mass limit of detection (MLOD). The CLOD relates to the concentration of the individual sample, whereas the MLOD describes what the instrument can measure. For example, the CLOD of many peptides is around 1 |ag/mL using absorption detection at 200 nm without stacking. If 10 nL of that material is injected and detected at three times the baseline noise, the MLOD is 10 pg.

Another way of considering MLOD is based on the volume of the detector window If a l-|Xg/mL peptide solution is continuously aspirated through a capillary, then for a 1-nL detector window, the amount of material in the window at any given time is 1 pg. Thus, the MLOD can be manipulated by selecting the size of the detection window. It is frequently possible to improve the MLOD at the expense of the CLOD by compressing the detection window. For most analytical problems, the CLOD is the more important parameter, since it relates to the minimum detectable quantity of a solute in the sample of interest. In extreme cases where the amount of available sample is minuscule, the MLOD becomes the more important parameter describing the LOD.

It is easily concluded the HPCE has excellent MLODs but poor CLODs, especially when compared with optical detection in HPLC. This is compensated for in part by the use of extended path length capillaries, online stacking, and offline trace enrichment.

9.4 DETECTION TECHNIQUES

A tabulation of detectors that have been used in HPCE is given in Table 9.1. Most of these detection schemes are not available on commercial systems or are not practical for everyday use. Commercial systems utilize absorption, fluorescence, laser-induced fluorescence, or conductivity detection. Interfacing to various forms of mass spectrometry is quite mature, though problems with sensitivity remain. Conductivity detection and indirect detection have been covered in Section 3.6.

9.4 Detection Techniques 369

Table 9.1 Detectors for HPCE

Technique References

Absorbance Absorbance, diode array Absorbance, extended pathlength Absorbance, indirect Absorbance, photothermal Capillary vibration Chemiluminescence Circular dichroism Conductivity Conductivity, indirect Conductivity, suppressed Concentration gradient Electrochemical, ampeometric Electrochemical, indirect Fluorescence Fluorescence, indirect Fluorescence, laser-induced Fluorescence, microscopy Fluorescence, multiwavelength Inductively coupled plasma Inductively coupled plasma mass spectrometry Ion mobility spectrometry Mass spectrometry, electrospray Mass spectrometry, fast atom bombardment Mass spectrometry, ion trap Mass spectrometry, magnetic sector Mass spectrometry, tandem Mass spectrometry, time-of-flight

NMR Oscillometric detection Phosphorescence, sensitized Potentiometric (ion-selective) detection Radioactivity Raman Reaction detector, affinity Reaction detector, postcapillary Refractive index

4-6 7-10 11-17 18-26 27-30 31-33 34-37 38 39-43 44 43,46 47-49 50-54 55 56-60 61-64 65-74 75-80 81-83 84-86 87-90 91 213,218-227 92-94 95-99 93,107, 108 100-106 97-98, 106 109-115 116-121 122 123 124-127 128-132 133,134 137-139 57, 135, 136 140-143

More than 150 papers have appeared reporting on electrochemical detection. This mode of detection is targeted primarily for catecholamine detection in single nerve cells, though many other applications are possible. It is unlikely that this mode will become commercially available in the near future, due to the difficulty of fabricating microelectrodes. That is unfortunate, since the LOD can


approach that of fluorescence for electroactive solutes. These detectors may become more practical with microfabricated devices (144-146).

Detectors fall into one of two broad categories:

1. Bulk property detectors 2. Solute property detectors

The bulk property detector measures a general property of matter. Refractive index and conductivity detection are the most important examples of the class. These detectors are not selective; they are universal. They are often less sensitive than solute property detectors. The sensitivity is enhanced if a solute's measured property is maximally differentiated from that of the BGE. This adds an additional constraint to methods development.

Solute property detectors measure a physical property that is specific to the solute as compared with the BGE. These are represented by absorption, fluorescence, electrochemical, and radioactivity detectors.

Fluorescence detection is far more sensitive and selective than absorption detection. All molecules that fluoresce must first absorb light. The converse is not true; most molecules do not fluoresce. The resultant selectivity is a double-edged sword. Since the technique is selective, derivatization is frequently required to take advantage of the detector's inherent high sensitivity. Sensitivity of detection is high, since a low signal level above a very dark background is more easily measured than is a small difference between two high-intensity signals, as in absorption detection. With the use of the laser as the excitation source, detection problems are virtually eliminated because of the extreme sensitivity. However, derivatization is often required, since most solutes do not have native fluorescence.

Information-rich detection is particularly important in HPCE. Since fraction collection is difficult, it is important to obtain additional information about a solute online. The mass spectrometer and multiwavelength absorption detectors are the most important examples of the group of detectors. Of the multi-wavelength detectors, the diode array detector in particular is very useful during methods development, since it can aid in peak tracking.

Another means of categorizing detectors is the nature of the response toward the solute. Most detectors used for HPCE are concentration-sensitive. They respond in proportion to the concentration of the solute as it traverses the detection window. The mass spectrometer is the most notable exception of the group. Sensitivity is based on the number of formed ions, so that this instrument responds to the mass of material that enters the source.

9.5 BAND BROADENING

Two detector-related features can contribute to band broadening: the width of the detector window and the detector time constant. For all practical purposes,

9.5 Band Broadening 371

the detector-related contribution to peak variance is the same as the injection contribution (147, 148), and so

< - ^ , (9.3)

where (j\^^ is the detector-related peak variance and l^ is the length of the detector window. Data given in Table 8.2 can be used to determine if the detector volume is sufficiently small. A 500,000-plate peak with t^ = 10 min requires I et < 0.56 mm. Most commercial absorption detectors meet this constraint.

All detectors employ smoothing algorithms to decrease detector noise. To the user, the adjustable parameter is the time constant or rise time. Virtually all modern detectors use rise-time functionality, normally contained within the system software. The rise time is approximately twice the effective RC time constant. These newer algorithms permit enhanced smoothing without excessive band broadening.

Insufficient detector time constant settings needlessly sacrifice signal-to-noise (S/N) ratio, whereas higher values can result in band broadening. The detector time constant should be no greater than 10-20% of the temporal peak variance. Table 9.2 provides a guide for selecting the time constant based on the temporal peak variance.

Figure 9.2 illustrates the impact of detector rise time for a 40,000-plate separation with Vjn = 2.1 mm/s. Note that 0.5 s is slightly higher than optimal, as evidenced by the reduced peak height (and the tabular value for (7 ), but S/N still is improved. Even at a rise time of 1 s, the peak is still Gaussian with further improvement in S/N. As with the situation with injection, it is possible to trade some plates for improved sensitivity For high-resolution separations, the trade-off is far more difficult.

Table 9.2 Selection of Detector Time Constant for a 100-cm Capillary and Mn = 2 mm/s

N

50,000

100,000

500,000

1,000,000

Gy (mm)^

4.47

3.16

1.41

1.00

CTt ( S ) ' '

2.24

1.58

0.70

0.50

Rise Time'

0.5

0.2

0.1

0.05

'a, = iL/NY .

^Converted into units of rise time and rounded to numbers consistent with commercial detector settings.


0.1

W^'-vrf'*^

FIGURE 9.2 Impact of detector rise time on signal, peak shape, and noise. Capillary: 46 cm (35 cm to detector) x 50 |Xm i.d.; buffer: 50 mM SDS, 20 mM borate, pH 9.3; voltage: 22 kV, 33 ^A; detection: UV, 220 nm; solute: diflunisal, 13.8 |Xg/mL; noise traces at left of peak run at chart speed of 1 cm/min, attenuation 4x; signal traces run chart speed of 20 cm/min, attenuation 8x. Rise-time values in seconds are to the left of each peak.

9.6 ABSORPTION DETECTION

A. TYPES OF DETECTORS

Ultraviolet/visible absorption detection is by far the most popular technique used today. Several types of absorption detectors are available on commercial instrumentation, including:

1. Fixed-wavelength detectors using mercury, zinc, or cadmium lamps, with wavelength selection by filters

2. Variable-wavelength detectors using a deuterium or tungsten lamp, with wavelength selection by monochromator

3. Filter photometers using a deuterium lamp, with wavelength selection by filters

4. Scanning UV detectors 5. Photodiode array detectors

Each of these absorption detectors has certain attributes that are useful in HPCE. Clearly, multiwavelength detectors such as the photodiode array and scanning UV detectors are valuable, since spectral as well as electrophoretic information can be displayed (Figure 9.3). The scanning UV detector is fre-

9.6 Absorption Detection 373

quently less sensitive when used in the scanning mode, since signal averaging must be carried out more rapidly than for single-wavelength detection. When using the photodiode array detector in conjunction with coated capillaries, beware of deterioration of the capillary coating. Since all wavelengths of light are passed through the capillary, the intense radiation may photodegrade the coating. For DNA applications where 260 nm is the wavelength of detection, a bandpass filter is available to eliminate this problem.

Even a filter photometer can be invaluable. Wavelength calibration never varies, and so the lines defined by the common atomic vapor lamps such as mercury, zinc, and cadmium can be considered primary standards. Another application is for low-UV detection. The simple optical design reduces the generation of UV-absorbing ozone, which causes problems with monochromator-based instruments. The use of the 185-nm mercury line becomes practical in CZE with certain buffers, since the short optical pathlength minimizes the background

^Hewlett-Packard, Wilmington, DE.

FLUORESCEIN (1 m g / m l )

Inj:

Ru

400 450 500 550

FIGURE 9.3 Diode array detection of fluorescein, 1 mg/mL. Capillary: 72 cm (50 cm to detector) X 50 l m i.d.; buffer: 20 mM CAPS, pH 11; injection: 10 kV, 10 s.


absorption. Since peptides absorb strongly at 185 nm, sensitivity is frequently enhanced, as shown in Figure 9.4.

B. SENSITIVITY OF DETECTION

The sensitivity of absorption detection can be calculated from the rearranged form of Beer's law equation:

CLOD = 5 X 10"

(a)(b) (5000)(5 x 10"^) = 2 X 10"^M, (9.4)

Where A = absorbance (AU), a = molar absorptivity (AU cm~^M"^), h = capillary diameter or optical pathlength (cm), and C = concentration (M). The noise of a good detector is typically 5 x 10"^ AU. A modest chromophore has a molar absorptivity of 5000. Then, in a 50-|im-i.d. capillary, a CLOD of 2 X 10"^ M is obtained at a signal-to-noise ratio of 1, assuming no other sources of band broadening.

—r— 14 6 10

Time (min.) FIGURE 9.4 Detection of impurities in a synthetic pentapeptide (Asp-Ser-Asp-Pro-Arg) at 214 and 185 nm. Capillary: 60 cm x 75 |im i.d.; buffer: 100 mM phosphoric acid, pH 2; injection: gravity, 10 cm, 10 s; voltage: (upper trace) 12 kV; 214 nm, 1 mg/mL; (lower trace) 12 kV; 185 nm, 335 jig/mL. Courtesy of Waters Associates.


Equation (9.4) is important, since it provides the basis for optimizing the sensitivity of absorption detection. Some of these features are within the control of the analyst. For example, the detector noise can be minimized by selecting BGE reagents that do not absorb at the wavelength of detection. The capillary pathlength can be increased by using extended pathlength capillaries. The molar absorptivity can be optimized by selecting the appropriate wavelength of detection, or through derivatization techniques. The sample concentration can also be increased by offline trace enrichment, or online using one of the stacking techniques.

C. SELECTING THE OPTIMAL WAVELENGTH OF DETECTION

Because of the short optical pathlength, the selection of the optimal wavelength is frequently different from that in HPLC. With a 1-cm pathlength, HPLC mobile phases absorb too strongly to use low-UV wavelengths. This is particularly troublesome with gradient elution, since substantial baseline drift is generally encountered. In HPCE with a variable-wavelength absorption detector, the optimal signal-to-noise ratio for peptides is found at 200 nm. This is illustrated in Figure 9.5 with electropherograms of some dynorphins obtained at wavelengths of 200, 214, and 280 nm.

To select the optimal wavelength, it is necessary to plot the signal-to-noise ratio. It is easy to do this in a few minutes without performing any real runs. First, determine the effective detector noise at 5-nm intervals using BGE and applied voltage. Similarly, aspirate some sample through the capillary, and repeat each measurement. It is generally necessary to re-autozero the instrument versus the BGE after each wavelength change. Finally, calculate the signal-to-noise ratio at each wavelength, and select the optimal value. The photodiode array detector makes this process especially simple.

D. INCREASING THE OPTICAL PATHLENGTH

Increasing the optical pathlength of the capillary window should increase S/N simply as a result of Beer's law (11). This may be achieved in several ways. One cell is commercially available^ and configured as shown in Figure 9.6. This so-called Z-cell has a pathlength of 3 mm. The optical window is integral to the capillary. This device is expensive, and if the capillary is fouled, it may have to be replaced. There has been some success with sleeving a new capillary onto the cell using some Teflon tubing to make the joint, and methods for joining capillaries have been reported as well (149).

2LC Packings, 80 Caroline Street, San Francisco, CA 94103. The cell fits the ABI 270A, ABI 785 detector, and Waters Quanta instruments.


s m

—-M»uyi yv* \ * ^

TIME (min.) FIGURE 9.5 Optimization of detector wavelength for peptide separations. Capillary: 65 cm (43 cm to detector) x 50 jim i.d.; buffer: 20 mM citrate, pH 2.5; voltage: 25 kV; temperature: 30°C; sample: dynorphins, 50 |J,g/mL. (A) 200 nm, attenuation 8x; (B) 214 nm, attenuation 4x; (C) 280 nm, attenuation 4x.

Electropherograms, given in Figure 9.7, showed an S/N improvement of only sixfold despite a 60-times-longer pathlength. Inadequate focusing of the light is responsible for this disproportionate observation, and when optimized, the sensitivity increases yet further (15). The sensitivity of absorption detectors is related to the amount of light reaching the photodiodes. Since the Z-cell attenuates the light, the S/N improvements are not proportional to the increased path length. The increased volume of the flowcell gives rise to 20-30% band broadening. Improved sensitivity will be found only when the UV background absorption of the BGE is low.

A high-sensitivity cell is also available from Hewlett-Packard, the configuration of which is shown in Figure 9.8. This cell comes in three parts: an inlet


UV light

FIGURE 9.6 Schematic of a 3-mm Z-shaped capillary flow cell. 1 = shim with centered 300-|lm-i.d. hole; 2 = plastic disks; 3 = fused-silica capillary. Redrawn with permission from J. Chromatogr., 542, 439 (1991), copyright © 1991 Elsevier Science Publishers.

capillary, an outlet capillary, and the cell body itself. The inlet and outlet capillaries are specially manufactured with one flared end each. The flared ends butt snugly into the cell body to reduce the dead volume. Improvements in sensitivity of an order of magnitude have been reported (17). The cell is only available for 75-|Llm-i.d. capillaries and provides a 1-mm path length. The proper assembly of the device takes some practice. Band broadening based on the


0.01 AU

3mm Z-Cell

ON-CAPILLARY k

I— 0.0 5.0 10.0

TIME (min)

34 J L L

15.0

FIGURE 9.7 Separation of nucleosides by MECC with the capillary Z-cell. Capillary: 60 cm (40 cm to detector) x 75 jim i.d.; buffer: 6 mM borate-10 mM phosphate, 75 mM SDS, pH 8.5; voltage: 11 kV; injection: gravity, 10 cm, 5 s; detection: Uy 254 nm. Solutes: 50 fXg/mL each: (1) 2'-deoxyxy-tidine; (2) 2'-deoxyguanosine; (3) 2'-deoxyguanosine-5'-monophosphate; (4) 2'-deoxycytidine-5'-monophosphate. Reprinted with permission from J. Chromatogr, 542,439 (1991), copyright © 1991 Elsevier Science Publishers.

increased optical path will occur with this cell, and so adequate resolution must be designed into the separation.

Another, simpler device for extending the capillary pathlength is known as the "bubble cell." This cell is integral with the capillary and is inexpensive. A photo of the bubble cell is shown in Figure 9.9. Several versions are available from Hewlett Packard including 25 |Lim, BF 5; 50 |Lim, BF 3; and 75 |Lim; BF 2.7. BF, the bubble factor, is the degree of enlargement of the capillary i.d. If the background absorbance of the BGE is low, the LOD decrease is close to the bubble factor. Of particular note is the 25-jLlm, BF 5 capillary This capillary provides the advantages of a narrow diameter, yet sensitivity does not suffer. By selecting the appropriate (proportionately narrower) optical slit, no band broadening is observed when using the bubble cell.

9.7 Fluorescence Detection 379

FIGURE 9.8 Drawing of a high-sensitivity cell. The inlet and outlet capillaries are separate. A 1-mm optical pathlength is obtained. Courtesy of Hewlett-Packard.

Sensitivity can be enhanced by viewing down the long axis of a rectangular capillary, but after the initial report (3), no further work has appeared. Square capillaries are also available from Polymicro Technologies. The optical surface should be superior to that of a round capillary, but no reports have appeared in the literature describing the advantages.

9.7 FLUORESCENCE DETECTION

A. BASIC CONCEPTS

Instrumentation for non-laser-based fluorescence detection is rare on commercial devices. With the Dionix instrument no longer available, the only unit than

FIGURE 9.9 Photo of a "bubble cell." Courtesy of Hewlett-Packard.


can be adapted for fluorescence detection is the ThermoQuest Crystal 300 modular system. Fluorescence detection, even with noncoherent light sources, can improve the limits of detection by several orders of magnitude over that of absorbance detection. The advantage of conventional light sources is tunabil-ity; lasers operate only at discrete wavelengths.

The fluorescence advantage is the result of detection against a very dark background. An optical transducer, the photomultiplier tube (PMT), is very sensitive to minuscule amounts of emitted light. Unlike absorbance detection, fluorescence detection is extremely dependent on the instrumental design. The fundamental equation governing fluorescence is

I, = 0,I,abcE^E,,E^E^^,, (9.5)

where If is the measured fluorescence intensity, ^f is the quantum yield (photons emitted/photons absorbed), I^ is the excitation power of the light source, a, b, and c are the Beer's law terms, and the E terms are the efficiencies of the excitation monochromator or filter, the flow cell, the emission monochromator or filter, and the PMT, respectively The situation is further obscured, since J and all of the efficiency terms show wavelength dependence. This means that instrumental parameters must be considered carefully when developing an assay. Clearly, the optimization scheme for fluorescence is more complicated than one for absorption detection, but the extra sensitivity and selectivity can be well worth the effort.

Various light sources are useful for fluorescence. Deuterium is useful for low-UV excitation, whereas the xenon arc is superior in the near-UV to visible region. Xenon lamps can be very powerful, and limits of detection by CZE of 2 ng/mL for fluorescein are possible with simple fiber-optic collection of fluorescence emission (57). This is a 15-fold improvement over absorbance detection v^th a tungsten lamp at 490 nm, and 100-fold improvement over deuterium-based absorption at 240 nm.

The design of a fluorescence detector for CE is relatively simple. One such design, which uses fiber optics to collect emitted radiation, is shown in Figure 9.10 (57). Many other ingenious designs are also possible (56, 59), including epiillumination microscopy (75, 76, 78).

While all compounds that fluoresce must absorb, the converse is not true, and so fluorescence can be selective. This is a double-edged sword, since the technique is amenable to fewer compounds. Often, derivatization is required. The selectivity of fluorescence is illustrated in Figure 9.11 with a peptide map (57). Only tryptophan-containing peptides have significant fluorescence.

B. OPTIMIZATION

Since the intensity of fluorescence is directly proportional to the lamp energy at the excitation wavelength, the proper wavelength must be selected. The opti-


MONOCHROMATOR

PHOTOMETER

FILTER (S)

FIGURE 9.10 Schematic of a tunable combination absorption-fluorescence detector for HPCE. Key: (A) slit; (OE) sapphire lens; (Fl, F2) optical fibers; (D) photodiode; (S) emission filter. This device is not commercially available. Reprinted with permission from Anal. Chem., 63,417 (1991), copyright © 1991 Am. Chem. Soc.

mal excitation wavelength is equal to the product of the lamp energy output and the strength of the absorption band. With a deuterium lamp for excitation, this wavelength frequently corresponds to absorption maxima of solutes in the low UV The xenon arc source is better suited to solutes that absorb in the near-UV to the visible spectral region. For the tungsten lamp, only absorption bands in the visible wavelength region are useful.

Having selected an excitation wavelength, the emission wavelengths must be selected. First, obtain a fluorescence emission scan of the solute using a scanning spectroflurometer. Cutoff or bandpass filters are usually employed to select emission wavelengths in HPCE. Bandpass filters with a 10- to 25-nm bandwidth are easiest to use, since they can be matched to the emission maximum. Interference from specular, Raman, and Rayleigh scattering is less likely with bandpass filters. If a monochromator is used for excitation, never select a bandpass filter that is a whole-number multiple of the excitation wavelength. The monochromator passes higher orders of wavelengths, which will raise the background. For example, if 250 nm is used for excitation, do not select a 500-nm bandpass filter.

It is useful to perform an emission scan of the BGE using the proposed excitation wavelength. Search the emission spectrum for the Raman band. The Raman band is redshifted 10-100 nm (or more) from the obvious Rayleigh band. This shift is excitation-wavelength- and solvent-dependent. Using 254-nm excitation.


Ay Absorbance: Deuterium at 200 nm

B, Fluorescence: Xenon with Xexs200, Em: 305LP

C, Fluorescence: Xenon with Xex=280, Em: 305LP

Buffer: 20 mM sodium phosphate 50 mM hexane sulfonic acid, pH 2.5

—T— 18

TIME (min)

FIGURE 9.11 Native fluorescence detection of tryptophan-containing peptides from a tryptic digest of -lactoglobulin. (A) absorbance detection at 200 nm; (B) fluorescence detection with deuterium lamp excitation, emission selected with a 305-nm-long wavepass filter; (C) fluorescence detection with xenon arc excitation, emission selected with a 305-nm-long wavepass filter. Capillary: 72 cm (50 cm to detector) x 50 |J.m i.d.; buffer: 20 mM sodium phosphate, 50 mM hexane sulfonic acid, pH 2.5; field strength: 278 V/cm; temperature: 30°C; injection: vacuum, 2 s; initial protein concentration: 20 nmol/mL. Reprinted with permission from Anal. Chem., 63, 417 (1991), copyright © 1991 Am. Chem. Soc.


the shift is approximately 20 nm; at 365-nm excitation, the shift reaches 60 nm; whereas at 546-nm, a Raman shift of 100 nm is normal. If the Raman band overlaps with your proposed emission filter, adjust the excitation wavelength or emission filter to eliminate the interference. Failure to do so can reduce detector sensitivity by an order of magnitude.

Any fluorescent impurities in the background electrolyte should be avoided as well. Paying careful attention to these details will optimize the limit of detection. Since excitation and emission bands are quite broad, there is considerable latitude for selecting the operating conditions. Maximizing the distance between excitation and emission, avoiding Raman bands, and optimizing wavelengths to maximal absorption and emission provide the best results. Optimal S/N can usually be obtained empirically after a few experiments.

C. LASER-INDUCED FLUORESCENCE ( L I F )

The high photon flux and spatial coherence (focusing capability) of laser light provide excellent properties for a fluorescence excitation source. Since laser lines are monochromatic, background elevations from Rayleigh and Raman scattering are easily avoided by selection of the appropriate emission wavelengths. Laser-induced fluorescence was first reported in 1988 (65), and the next 10 years produced more than 600 papers in the field. A 1997 review article traces the history and direction of this technique (68). For details on bioapplications, especially DNA, reference (150) should be consulted. A wide variety of lasers can be used for LIF detection, as shown in Table 9.3. The low-UV lasers, which are useful for measuring native fluorescence of small molecules, are not compatible with the Beckman instrument unless a UV-transparent fiber optic is employed. The krypton-fluoride laser emits at 248 nm and can be used in conjunction with homemade systems (151, 152). Some of these other lasers are large, expensive, or unreliable. The most useful lasers for LIF detection are the argon-ion, helium-cadmium, helium-neon, and diode lasers.

Low-power argon-ion lasers address cost and ruggedness issues effectively. Very clever designs to reduce scattering have been described in the literature. These include complex flowcells that use sheath fluids that are refractive index matched to the run buffer (153-155). Changes in the refractive index between abutting surfaces is the root cause of light scattering. In the absence of scattering and with picoliter-volume flowcells, MLODs approach single-molecule detection. Simple designs such as that of Drossman et al. (156), shown in Figure 9.12, can provide LODs of 10"^^ M for fluorescein. This corresponds to 60,000 molecules, a 10-nL injection of a 10~ ^ M solution.

Commercial LIF instrumentation is available from Beckman as an accessory to the P \ACE and MDQ instruments. The LIF detector includes a power supply, 3-mW argon-ion laser, and a unique device to collect emitted fluorescence


Table 9.3 Laser Light Sources for LIF Detection

Laser

Ar-ion (air-cooled)

Ar-ion (full frame)

Ar-ion (full frame, frequency-doubled)

ArKr

HeNe

Excimer KrF (pulsed) XeCl (pulsed)

Nitrogen (pulsed)

Nitrogen pumped dye (tunable)

Solid state YAG (frequency-doubled) YAG (frequency-quadrupled)

Diode lasers Frequency-doubled (LiNb03) Frequency-doubled (KTP) Frequency-tripled (Nd-doped YLiF)

Available Wavelengths (nm)

457,472,476,488,496,501,

275,300,305,333,351,364, 476,488,496,501,514

229,238,244, 248, 257

350-360, 457, 472, 476, 488, 521,514,521,531,568,647,

543, 594, 604, 612, 633

248 308

337

360-950

532 266

415 424 349

514

385, 457, 472,

496, 501, 514, 752

Data from reference (150).

(Figure 9.13). Laser light is transmitted to the capillary via a fiber-optic cable. Fluorescence is collected using an ellipsoidal mirror and focused backward toward the photomultipHer tube. A center hole in the flowcell allows most of the laser light to be directed away from the PMT, and a filter removes the balance. The company reported an LOD for fluorescein of 10" ^ M. The PE Biosystems 3700 DNA Sequencer employs an argon-ion laser with the beam transmitted perpendicularly through the entire 96-capillary array. The effluent from the capillaries is mixed with a sheath flow fluid, and the ensuing fluorescence is imaged on a charged-coupled device. While detection is postcapillary, the bands are still broadened as described in Section 9.1, since slower moving (later eluting) solutes receive more postcapillary dilution. Without LIF detection, the field of DNA sequencing would not have advanced so quickly.

Semiconductor lasers may develop as an alternative laser light source. The advantages herein are based on size, cost, and stability. At present, these lasers are available in quantity only for visible and near-lR wavelengths. Frequency doubling is required to reach lower wavelengths. One such laser, a 2.5-mW, 635-nm diode laser, is available on the Beckman LIF detector.

9.8 Derivatization 385

ARGON XON

LASER

I

I

BEAM EXPANDER

BPf

CAP

SP

BPF

PMT

FIGURE 9.12 Schematic of a simple laser fluorescence detection system for CE. Redrawn with permission from Anal Chem., 62, 900 (1990), copyright © 1990 Am. Chem. Soc.

Capillary Tube

Fiber-optic cable

Ellipsoidal Mirror

Beam-Block

FIGURE 9.13 Beckman Instruments LIF detector schematic. Courtesy of Beckman Instruments.

9.8 DERIVATIZATION

Since few molecules have native fluorescence, derivatization is frequently required. Likewise, when a solute is a poor chromophore, derivatization should

3 8 6 Chapter 9 Detection

be considered. Both pre- and postcapillary derivatization can be employed to enhance CE detection. Precapillary derivatization can be performed on current instrumentation, since no instrumental adaptations are required. Virtually all of the derivatizing agents used for liquid chromatography can be employed in CE. The subject has been reviewed for all forms of HPCE derivatization including that for Uy for fluorescence, and for LIE detection (157, 158). Hundreds of papers have appeared on this subject. The pros and cons of derivatization were covered in Section 4.91.

Table 9.4 contains a sampling of some commonly used reagents, along with conditions for fluorescence detection. If LIE detection is employed, the tag selection is further dictated by the available laser wavelengths. Since the argon-ion laser is the most commonly used, tags have been developed that absorb at 488 nm, the wavelength where this laser emits. The tags have also been designed to fluoresce at wavelengths away from Raman scattering. Many of these tags or dyes are available from Molecular Probes (Eugene, OR). Table 9.5 presents a selection of reagents, along with the mode of HPCE, the detection technique, and the solutes studied.

One of the first reagents specifically designed for HPCE with LIE is 3-(4-car-boxylbenzoyl)-2-quinolinecarboxaldehyde (CBQCA). Reacting with primary

Table 9.4 Fluorescent Derivatization Reagents

Reagent

a-Dansyl chloride^

NBD chloride

Fluorescamine

o-Phthalaldehyde

Dansyl hydrazine

Naphthalenedialdehyde

CBQCA'

Fluorescein Isothiocyanate

Bromomethylcoumarin

Thiazole Orange

APTS^

Reacts with

1°, 2° amines, phenols

1°, 2° amines

1° amines

1° amines

Aldehydes and ketones

1° amines

1° amines

1° amines

Carboxyhc acids

DNA intercalator

Carbohydrates

Excitation X (nm)^

360

420

390

350

340

442

442

488

325

488

455 (488 with laser)

Emission X (nm)''

520

540

475

440

525

490

550

525

430

520

512

^Emission and excitation wavelengths are solute and solvent dependent. A number of the excitation wavelengths correspond to laser hnes (325, 442, and 488 nm) and do not correspond to the actual excitation maxima. ^The emission wavelength is frequently selected to avoid the Raman band and may not correspond to the actual emission maximum. ^Dansyl chloride has a poor quantum yield in water. '^3-(4-carboxybenzoyl)-2-quinolinecarboxaldehyde. ^9-Aminopyrene-l ,4,6-trisulfonic acid.


Table 9.5 Reagents for Precapillary Derivatization

Reagent

AEC

APIS

BPB

CBQCA

Dansyl chloride

Dansyl hydrazine

FITC

FLEC

Fluorescamine

Fluorescein

FMOC

NBD-fluoride

NDA

NTAA

OPA

PA

PITC

CE Mode

MECC

CZE CZE

CZE

MECC MECC MECC

MECC

CZE

CZE CZE MECC

MECC MECC

CZE MECC

MECC

CZE/MECC

CZE MECC

CZE CZE PN

MECC

CZE MECC

CZE

CZE

Detection

UV/LIF

LIF LIF

UV

LIE LIE LIF

F

LIE

LIE LIE F

UV UV/LIF

LIE F

LIF

F/LIF

LIF LIF

LIF LIF/F UV/LIF

UV

LIF UV/F

UV

UV

Solute

Amino acids

Oligosaccharides Disaccharides

Organic acids

Amino acids Peptides Amino sugars

Amino acids

Sugars

Amino acids Aliphatic amines Polyamines

Amino acid enantiomers Amino acid enantiomers

Marine toxins Polyamines

Phenoxyacid herbicides

Amino acids

Amino acid enantiomers Dipeptides

Amino acids Jeffamine oligomers SDS-proteins

Organo Pb, Hg compounds

Amino acids Amino acid enantiomers

Nonionic surfactants

Maillard products

Reference

159

160 161

162

163 164 165

167

166

70 168 169

170 171

172 173

174

57

175 176

177 178 179

180

177 181

183

182

AEC = 2- (9-anthryl) ethyl chloroformate; APTS = 9-aminopyrene-l,4,6-trisulfonic acid; BPB = bro-mophenacyl bromide; CBQCA = 3-4-carboxybenzoyl-2-quinolinecarboxaldehyde; FITC = fluorescein isothiocyanate; FLEC = 9-fluorenyl ethyl chloroformate; FMOC = 9-fluoroenylmethyl chloroformate; NDA = naphthene-2,3-dicarboxaldehyde; NDB = 4-fluoro-7-nitro-2,l,3-benzo-xadiazole; NTAA = nitrilotriacetic acid; OPA = O-phthalaldehyde; PA = phthalic anhydride; PITC = phenylisothiocyanate.

amines, the absorption maxima of this tag match well with the 442-nm line of the helium-cadmium laser. This reagent produces MLODs down to 10" ^ M, as illustrated in Figure 9.14 for a tryptic digest of 1.9 pg lysozyme.

When using a nonlaser source or a KrF laser, FMOC (2-fluorenylmethyl-chloroformate) is ideal for derivatizing primary and secondary amines (57,184). Rapid derivatization, stable products, and high sensitivity are features of this


10 15 Time (min)

20 25 30

FIGURE 9.14 Electropheorgram of CBQCA-derivatized amino acids from a sample representing 1.9 pg hydrolyzed lysozyme. Capillary: 100 cm (70 cm to detector) x 50 |lm i.d.; buffer: 50 mM TES, 50 mM SDS, pH 7.02; voltage: 25 kV (14 |LiA); detection: He-Cd laser, 442 nm. Key: (1) Arg; (2) Trp; (3) Tyr; (4) His; (5) Met; (6) He; (7) Gin; (8) Asn; (9) Thr; (10) Phe; (11) Leu; (12) Val; (13) Ser; (14) Ala; (15) Gly; (16) Glu; (17) Asp. Reprinted with permission from Anal. Chan., 63, 408 (1991), copyright © 1991 Am. Chem. Soc.

reagent. In conjunction with a chiral recognition additive, enantiomeric separations are possible using this tag. This solves most sensitivity issues with regard to the trace enantiomer, particularly if LIF detection is used.

An achiral separation of some amino acids by MECC is shown in Figure 9.15 (57). FMOC is an ideal derivatizing reagent. Likewise, the chiral derivatizing reagent FLEC (2-fluorenylethyl-N-chloroformate) is a superior reagent. In this case, MECC with an achiral surfactant is used to resolve the diastereom'ers.

The DNA-sequencing reaction chemistry is designed to provide a bound fluorescent tag as well. All DNA sequencers use LIF detection. Differences in the wavelengths of emission from each of four different dyes are used to assign the sequence of DNA bases.

< >

lij X

a.

a

o DC

3

i t

-T 2 0 6 8 10 12 14

TIME (ifiiti) FIGURE 9.15 Separation of FMOC-derivatized amino acids. Capillary: 62 cm (40 cm to detector) X 50 |im i.d.; buffer: 20 mM borate, pH 9.5, 25 mM SDS; field strength: 416 V/cm; temperature: 30°C; detection: xenon arc fluorescence; excitation: 260 nm; emission: 305-nm-long wavepass filter. The CLOD is 10 ng/mL. Reprinted with permission from And. Chem., 263, 417 (1991), copyright © 1991 Am. Chem. Soc.

One can consider immunoassay a form of detection or even a biological derivatization (150, 185-190). There are basically two forms: competitive and direct (or noncompetitive) immunoassay. When a fluorescent-labeled antibody is available, the preferable noncompetitive immunoassay can be performed. Antibody and antigen are incubated together for a predetermined period of time, after which the free and bound forms are separated. Quantitation is either by the appearance of the immune complex or the disappearance of the labeled antibody In the competitive immunoassay, tagged antigen is mixed with untagged antibody and sample containing untagged antigen. The higher the concentration of


untagged antigen, the lower the concentration of the tagged antigen-antibody complex. The technique yields nonlinear calibration curves and should only be used when tagged antibody is not available.

LIF is particularly useful for detection, since the high sensitivity reduces the requirement for expensive labeled antibodies or antigens. The high specificity of fluorescence also reduces the potential for interference. If an antigen has native fluorescence, then a derivatized antibody is not required.

DNA intercalators have revolutionized the fields of genetic analysis and human identification. While not a derivatization chemistry, these reagents are added to the BGE. The noncovalent interaction between the dye and DNA modifies mobility, and so this is in effect a form of secondary equilibrium. It can also be thought of as a noncovalent on-capillary derivatization technique. More important is the impact of the dye on detection. When the dye is not bound to DNA, its fluorescence is quenched via interaction (collision) with water. Once bound to DNA, fluorescence quenching is reduced, and the fluorescence quantum yields increase dramatically The classical DNA intercalator, ethidium bromide, is seldom used today, since its wavelength of absorption does not match the emission wavelength of the argon-ion laser. Dyes such as TOTO, YOYO, TO-PRO, and YO-PRO are usually selected because of their favorable wavelength of absorption. These applications are covered in more detail in Chapter 6.

True on-capillary derivatization is possible as well. In this case, the front end of the capillary is used for the chemical reaction. As in the case of postcapillary reactions, the reaction kinetics need to be relatively rapid to minimize band broadening. This technique is ideal when sample volumes are extremely limited, as in the case of analysis of chemicals from a single cell (191).

On-capillary chemistry can also be employed for enzyme assays (192-194). Known as enzyme-mediated microanalysis (EMMA), the technique employs reaction between an enzyme and a substrate. Given an excess of substrate, it is possible to amplify a reaction product to provide very high sensitivity. In another variant of on-column chemistry, PCR and subsequent size separations of amplified DNA have been integrated using microfabricated devices (195, 196).

Postcapillary derivatization also works well with HPCE (57, 135, 155, 158, 197-199). This technique is less common than precapillary derivatization because of the lack of commercial equipment. The advantages of this approach are minimal sample handling and the ability to work with derivatives that have limited stability

The principal requirements for postcapillary derivatization are as follows:

1. The derivatizing reagent is invisible to the detector. 2. Rapid reactions occur. 3. The reactor provides minimal band broadening.

One such design is shown in Figure 9.16 (57). The basis for the function of this reactor is differential EOF The gap junction of this reactor is about 50 jam. With a separating capillary of 50 jim and a reactor capillary of 75 |lm, the


reagent is drawn into the reactor by differential EOF, since the volumetric requirements of the larger diameter capillary are not being fulfilled. Mixing is accomplished by convection.

OPA (o-phthalaldehyde) is an ideal postcapillary reagent. The reagent does not fluoresce, it is stable, and it reacts quickly with primary amines. The optimal conditions are 3.7 mM OPA in run buffer, 0.5% mercaptoethanol, 2% methanol, and 40°C (57). The CLOD for OPA glycine is 60 ng/mL with xenon arc fluorescence, X = 350 nm, M > 400 nm. The run-to-run reproducibility was about 1% using peak areas. The sensitivity of peptide mapping can be greatly enhanced relative to absorption detection with the PCRS system, as shown in Figure 9.17 (57). The concentration of the analytes in the absorbance electro-pherogram was 40 times greater than in the fluorescence run.

Among the other notable postcolumn reaction detectors is chemilumines-cence detection (34, 36, 37, 200, 201). Chemistries such as peroxyoxylate, acri-dinium, luminol, and firefly luciferase have all been reported. Elimination of

Fluorescence Cell Reactor Ceil

to w a ^ reservoir

Exploded View of Buffer Junction

/ 50 \m gap i

l/t6x.007teftontut)«

FIGURE 9.16 Schematic of a liquid junction postcapillary reaction system. Reprinted with permission from Anal. Chem., 63, 417 (1991), copyright © 1991 Am. Chem. Soc.


s^

"T" 0 2

"T" 8 10 12 14

TIME (min)

FIGURE 9.17 CZE of a tryptic digest of/3-lactoglobulin with UV detection at 200 nm (top) and postcapillary derivatization with OPA (bottom). Capillary: 62 cm (40 cm to detector) x 50 |Lim i.d.; buffer: 20 mM borate, pH 9.5; field strength: 278 V/cm; postcapillary detection: xenon arc fluorescence; excitation: 390 nm; emission: 450-nm-long wavepass filter; sample concentration: absorption, 20 nmol/mL, fluorescence, 0,5 nmol/mL; injection: absorption, vacuum, 1 s, fluorescence, electrokinetic, 7 s at 5 kV Reprinted with permission from Anal. Chem., 63, 417 (1991), copyright © 1991 Am. Chem. Soc.

the excitation light source greatly diminishes the background light. As a result, the photomultiplier tube can be run at very high voltages, providing impressive sensitivity. Laserlike LODs on inexpensive instrumentation are possible; however, the simplicity of LIF and the lack of commercial postcapillary apparatus greatly limits application of this technique.

9.9 Mass Spectrometry 393

9.9 MASS SPECTROMETRY

A. INTRODUCTION

Coupling of HPCE to mass spectrometry (MS) is developing rapidly since first reported by Olivares et al. in 1987 (202). Looking back, the actual interfacing turned out not to be difficult, at least for electrospray ionization (ESI). Among the challenges were:

1. Providing an electrical contact in the absence of an outlet reservoir 2. Generation of sufficient fluid flow to maintain a stable spray 3. Finding compatible buffers and additives that are volatile and do not

raise the ion currents 4. Injecting a sufficient quantity of material to ensure detectability 5. Compatibility of the speed of the separation with the scan speed of the

mass spectrometer

There are two compatible ionization techniques: electrospray and fast-atom bombardment (FAB). Virtually all work has been reported using electrospray techniques. Both techniques generally require makeup flows to elevate the total flow rate to 1-10 |iL/min, although nanoelectrospray operates without a makeup flow. A syringe pump is typically used for reagent delivery to avoid the pulsations characteristic of reciprocating pumps, particularly at low backpressures and low flow rates. Since detection is performed postcapillary and most of flow is provided from the makeup solution, the problems with peak area normalization described in Section 9.1 are eliminated.

The HPCE instrumentation employed in mass spectrometry has several considerations.

1. The injection and capillary-filling mechanism must be pressure- rather than vacuum-driven. It is hard to imagine a simple means of connecting vacuum-driven equipment to any of the interfaces.

2. A design to safely route the capillary outside of the system is required. One such modification is shown for the Hewlett-Packard instrument in Figure 9.18.

3. To prevent siphoning, a lab-jack may be used to level the capillary inlet with the mass spectrometer.

4. The power supply should be capable of providing polarity switching and, in some cases, a negative electrode held at a potential other than ground. When the HP instrument is interfaced to an HP mass spectrometer (5989B MS Engine or 1100 LC/MSD), the negative electrode is grounded along with the electrospray. They must be linked with a grounding cable to be sure the are at the same ground.

Important: There are several different power supply configurations that can be used to couple HPCE instruments with mass spectrometers. It is critical that

Cassette

/

1 P^ 1 pi; '

==1 = ^ <v Mi-*ii-*L.*«^ tr:

) C3

HP 3DCE instrument Minimum total length = 83 cm

Minimum effective ler^th = 22 cm

Minimum total length without DAD = 55 cm

FIGURE 9.18 Illustration of a mass spectrometry capillary cartridge. Courtesy of Hewlett-Packard.

the appropriate hookups are employed, or power supphes may be damaged. In cases where the electrospray is at high voUage, a resistor sink must be used to prevent electrospray power supply damage. There may be current limitations as well. It is important to coordinate between vendors whenever using equipment from different manufacturers.

For most applications, it is useful to employ the on-board UV detector in conjunction with the MS interface. It is important to minimize the length of capillary between the detector and the interface; otherwise, the run time will be unnecessarily prolonged. The capillary length is dictated by the configuration of both the CE instrumentation and the interface. An extra 20-30 cm of capillary is normally used to make this connection. Wider bore capillaries of 75- to 100-|Lim i.d. are frequently used to increase the loading capacity of the system. When these capillaries are used, the system must be carefully leveled to prevent siphoning.

Most capillary electrophoretic techniques have been coupled to the mass spectrometer. These include CZE, CITP, CIEF, CEC, and CGE. The interface of MECC to the mass spectrometer has been more difficult because of interference from the surfactants. Partial-fill techniques have been used here, where the surfactant migrates back toward the inlet side of the capillary and does not enter the mass spectrometer. Table 9.6 contains references for interfacing these various techniques.


The coupling of CIEF with MS is most notable, since the data analogous to a two-dimensional electrophoretic map are generated in 30 min as opposed to two days. However, the sensitivity of the mass spectrometer is less than that of silver staining.

There has been only one report of coupling a gel-filled capillary to MS. Garcia and Henion (208) interfaced CGE using the lonSpray interface in 1992. They found that the high concentrations of urea typically used here do not elute from the gel. Common buffer systems such as Tris-borate-EDTA are retained as well. The negative-ion mode was employed to separate and monitor benzene and naphthalene sulfonates, dansyl amino acids, and polyacrylic acids.

The coupling of CEC to the mass spectrometer is advantageous compared with that of MECC, since surfactants are not used. The hydroorganic solvents used in CEC along with volatile buffers make interfacing relatively easy The use of CITP is particularly important because of the potential for a 500-fold trace enrichment of minor components in a mixture.

B. ELECTROSPRAY IONIZATION

The electrospray interface (ESI) solves many of the aforementioned challenges in coupling HPCE to the mass spectrometer. In this technique, ionization takes place at atmospheric pressure outside of the MS. The BGE should contain only volatile buffers and additives to avoid contamination of the MS by nonvolatile salts. Even when volatile additives are used, they can increase the chemical noise of the system. This is more important if a sheathless system is used.

Sometimes, the electrospray is grounded. If the electrospray is not grounded and is maintained at -H4 kV, then 26 kV is available for the CE separation. A variant of "pure" electrospray named lonSpray uses a nebulization gas to further assist spray formation (235). Thermal assists have also been used, but for flow rates that are far more rapid than ever encountered in HPCE (235).

Table 9.6 Interfacing of Various HPCE Techniques to the Mass Spectrometer

Technique References

CEC

CGE (size separations)

CIEF

CITP

CZE

MECC

203-207

208

96, 209-212

213-217

218-227, hundreds more

228-234


The electrospray generates small droplets, which begin to evaporate. Low surface tension facilitates droplet formation; otherwise, pneumatic nebulization may be required. High voltage is required to enhance the charging of the droplets. As evaporation continues, charged ions are brought into close proximity of each other. At some point, columbic explosions occur, and these take place many times, producing ever smaller droplets and finally gas-phase ions. These processes are illustrated in Figure 9.19.

While prodigious numbers of ions are produced by ESI, only a few find their way into the mass spectrometer. Despite the poor sampling efficiency, the sensitivity of ESI exceeds electron impact ionization by several orders of magnitude for ionic compounds. For nonpolar neutral compounds, ESI sensitivity can be poor and require collision-induced dissociation (CID) to obtain measurable signals, though this has not been reported for HPCE-ESI-MS.

Positively charged ions are produced through complex reactions between the solute and highly reactive ion clusters of water. Negative ions can be produced as well via reactions with O2", O", or complex hydrate clusters (235). Nitrogen oxides may also play a role in negative-ion formation. The selection of positive or negative ions is dictated by the polarity of the electrospray voltage, the pH of the buffer, and the setup of the mass spectrometer. Most CE-electrospray work has been done in the positive-ion mode, though the negative-ion mode should prove useful for many acidic substances. The electrospray is set at a voltage of about -5 kV for negative-ion production.

Elevation of the sample side of a 50-|im-i.d. capillary was reported to improve S/N, presumably through generation of a modest siphon-produced flow (219). It will probably be necessary to reduce the height differential should larger capillaries be used. A separation comparing CZE-UV with CZE-MS for some myo-

Nebuiizing gas

Slwath iiqiild

•ht+

Taylor cone

-+ " ^ ^ -" +''J> +t-."i %J^ iffC+ +U ++:f

SS tubos for sheatii liquid and aoballzing gas

CE capillary

FIGURE 9.19 The process of ion formation using the electrospray. Courtesy of Hewlett-Packard.


globins in shown in Figure 9.20. The migration times are somewhat longer by CZE-MS because of the extra capillary length that was used. Slightly better resolution and no evidence of significant band broadening was found, despite the production of hydrodynamic flow from capillary elevation.

Selection of the buffer and the makeup liquid is important. For positive-ion formation, 0.2% formic acid, 15 mM ammonium acetate, or 15 mM ammonium formate are useful volatile buffers. Lower buffer concentrations may be insufficient for separation; higher concentrations may lead to a reduction in the total ion current (236). Organic modifiers such as 5-25% methanol increase the ion response; methanol is superior to acetonitrile in this regard. Using more than 25% methanol caused bubble formation (236); however, higher concentrations of acetonitrile are possible. In addition, the speed of separation is faster using acetonitrile, since linear alcohols decrease the EOF Other acceptable reagents for both the BGE and sheath flow solution are trifluoroacetic acid and acetic acid.

As the solvent evaporates, ions are ejected directly into the gas phase and sampled into the vacuum region of the mass spectrometer through a series of

1§ 10

Time (min)

FIGURE 9.20 Comparison of CZE-UV (214 nm) and CZE-MS (total ion electropherogram) for a mixture of whale (MW 17,199.1), horse (MW 16950.7), and sheep (MW 16923.3) myoglobins. Buffer: 10 mM Tris, pH 8.3; field strength: 120 V/cm; injection size: 100 fmol. Reprinted with permission from J. Chromatogr., 559, 197 (1991), copyright © 1991 Elsevier Science Publishers.

skimmers. A potential difference of 3 kV between the electrospray and ion-sampling orifice facilitates sampling (236). The spray is best sampled at oblique angles to the MS inlet to reduce the background from the production of cluster ions (237). Nitrogen purge at the atmospheric side of the skimmer further reduces cluster-ion formation (236). Since ions are produced by the electro-spray, it is unnecessary to produce chemical or electron impact ionization inside of the instrument. Molecular ions form the predominant species. Fragmentation patterns can be produced and detected with a triple-quadrupole instrument via introduction of a collision gas such as argon at the second quadrupole (236).

Nearly all CE-electrospray work has been done using single- or triple-quadrupole instruments, though work has appeared using ion traps (96, 225, 238), time-of-flight instruments (109, 110, 239), and magnetic sector instruments (107). The quadrupole instruments have mass ranges of about 2400 Da. This is an miz ratio, where m is the mass and z is the charge. Electrospray produces multiply charged ions for proteins and oligonucleotides, and so it is possible to detect solutes with molecular weights over 100 kDa. The electrospray mass spectrum of myoglobin is shown in Figure 9.21.

The actual mass is determined as follows (235). For adjacent peaks, assume n2 = ni + 1, where n = number of charges. The detected mass (m^) is given by

m, = M + ni

(9.6)

where M is the actual mass and n^ is the number of charges. The measured m/z is the sum of the mass and the mass of the protons forming the positive ion. Then,

nil - 1 (9.7)

m.

100 942.7

\ 998.0 893.1

1063.3 1130.9 <W+13H)

1211.8 I

1304.8

13+

1 1 ^U4

^b*iirf m/z 1100 1200 1300

FIGURE 9.21 Electrospray MS of myoglobin. Reprinted with permission from The API Book, 1990, Sciex.


and

M = rijinij - 1 ) . (9.8)

The use of these equations produces a raw Uj value, which is rounded to the nearest integer. The integral n value is used to calculate the actual mass of solute as given in Table 9.7 for myoglobin. Production of ions in the vapor phase from molecules with very high masses is remarkable, as is the ability to detect and measure them on a quadrupole instrument with a limited mass range. Smith et a\. (240) reported on the determination of about 100 different peptides and proteins covering a mass range from 409 to 133,000. Most of these materials will be separable by CZE.

To overcome wall effects, Thibault et al. (241) employed a cationic surfactant coating along with charge reversal for some protein separations. Wall interactions were overcome, and good separations and spectra were obtained. It is expected that this approach, along with the use of other forms of capillary treatments, will permit separation and detection of most macromolecules.

One problem with multiple charges on a solute is loss of sensitivity that depends on molecular weight (242). These data are presented in Table 9.8. There are concentration-dependent issues as well. At lower concentrations, the relative peak intensities shift to higher charge states. The lower charge states include a series of adduct bands (data not shown) on the high-m/:^ side of each molecular ion. Despite these problems, high-sensitivity measurements in the

Table 9.7 Calculation of Molecular Weight of Myoglobin from Multiple-Charge Electrospray MS Data

^ 2

1304.8

1211.8

1130.9

1060.3

998.0

942.7

893.1

848.5

808.1

771.4

mi

1211.8

1130.9

1060.3

998.0

942.7

893.1

848.5

808.1

771.4

737.7

Raw Ui

13.03

13.97

15.00

16.00

17.03

17.99

19.00

19.98

20.99

21.86

^2

13

14

15

16

17

18

19

20

21

22

Average

Std dev.

% RSD

M

16,949.4

16,951.2

16,948.5

16,948.5

16,949.0

16,950.6

16,949.5

16,950.0

16,949.1

16,949.8

16,949.5

0.9

0.005

Reprinted with permission from The A?I Book, 1990, Sciex.

Table 9.8 Dependence of Signal Intensity on Molecular Mass

Mass

1,000

10,000

40,000

100,000

200,000

Ave. No . of Charges^

1

10

40

100

200

Peak Width {mJz)^

1

1

3

6

>6^

Ion Current (ions/s)'^

1 X 10^2

2 x 1 0 1 0

4x10^

3x10^

8x10^^

Âssumed that the average number of charges increases linearly vîth Mr and the distribution is centered on mJz 1000. ''Peak width due to microheterogeneity typical of large biopolymers and contributions of impurities, solvent adducts, etc. ' ESl production before sampling losses assuming an 80% ionization efficiency. Detected ion intensities are 4-5 orders of magnitude lovêr due to inefficiencies arising from sampling, transmission, and detection. ' Peak width of 6 m/z units is too large for individual charge states to be resolved; a peak width of <4 is required. ^For a peak width of 6 m/z units. Data from J. Chromatogr., 1990; 516:157.

full-scan mode are possible, as shown in Figure 9.22 for a scan of 10"^ M (23 fmol injected) cytochrome c (242). In practice, the sensitivity will be lower in the full-scan mode, since 90-s scans are impractical. In the single-ion-monitoring mode, the LOD of MS detection for peptides can be an order of magnitude lower than for UV detection.

The problem generated by the slow scan speed of the quadrupole is solved by employing time-of-flight (TOF) mass spectrometry. This is particularly important when high-speed separations are employed. Peaks can be missed or badly broadened using slow-scanning instruments. This problem is solved using the rapid-scanning TOF instrument (110).

There are four basic interface designs. The review article by Banks was used in part to help organize the following subsections (243):

1. Sheath flow 2. Sheathless flow (nanoelectrospray) 3. Liquid junction 4. Direct electrode

1. Sheath Flow

The electrospray interface designed by Smith and co-workers (219, 244) is illustrated in Figure 9.23. Sheath flow electrospray is the most common form of CE-MS, because it allows conventional capillaries to be used. The capillary is inserted into a tube, which is designed to deliver a sheath liquid around the cap-


700 23fmol

t ? ^ iSo 1300

FIGURE 9.22 ESI-MS of 1.5 x 10~ M cytochrome c obtained by direct infusion at 1 |alVmin during a 90-s acquisition period. Reprinted from J. Chromatogr., 516, 157 (1990), copyright © 1990 Elsevier Science Publishers.

illary outlet. A sheath gas may also be added, as shown in the figure. In modern designs, a metal sheath fluid tube is used to complete the electric circuit and provide the voltage for charging the electrospray droplets.

The sheath fluid provides both the electrical contact and the bulk of the Uquid for the electrospray. The flow from the capillary is perhaps 100 nlVmin, and the sheath flow is about 5 jLiIVmin. The sheath fluid must be sufficiently conductive to complete the circuit, but not so high in ionic strength to create arcing (245). The fluid must also have a low surface tension. Typically methanol-water-acetic acid mixtures meet these requirements. Even 100% methanol is sufficiently conductive and is preferred due to the reduction of chemical noise in the system (245).

In CE-ESI-MS of amphetamines (246), the best results were found with a sheath fluid of isopropanohwater, 50:50, with 0.5% formic acid. Acetonitrile, an aprotic solvent, was quite poor in this regard. The BGE was 100 mM formic acid. The system was optimized to maximize the sensitivity, reduce fragmentation, and minimize the dilution effect of the electrospray. Among the variables to be optimized are (the optimal values for the amphetamines are shown):

1. Sheath fluid makeup (50:50 lPA:water with 0.5% formic acid) 2. Sheath fluid flow rate (4 jLiL/min) 3. Electrospray voltage (3000 V) 4. Drying gas flow rate (6 L/min) 5. Skimmer voltage (50 V) 6. Temperature (200°C)


REMOTE ELECTRODE

CAPILLARY COLUMN

GLASS OUTER CAPILLARY

FIGURE 9.23 Schematic illustration of the electrospray interface for CE-MS. Reprinted with permission from J. Chromatogr., 559, 197 (1991), copyright 1991 Elsevier Science Publishers.

2. Sheathless System

In the sheathless system, the capillary outlet tip is drawn to an extremely fine point and coated with a metal to provide for electrical continuity (238, 247-249). Since no makeup fluid is required, the HPCE effluent is not diluted, which enhances sensitivity An insulating sheath gas can be used to prevent arcing from the high field strength at the narrow tip. Because there is no sheath fluid, there are more constraints on the buffer systems. It is essential that volatile buffers with low surface tension and low conductivity be used in this system. Since specialized capillaries and instrumentation are required for sheathless CE-ESI-MS, this mode has been used less frequently.

3. Liquid Junction Interface

The liquid junction interface is essentially a mixing "T" where the makeup fluid is added (250). A nebulizing gas is also used here. When pneumatically assisted electrospray is employed, the liquid junction can be used without actually pumping a makeup reagent (236). In this case, the nebulization gas provides a slight vacuum, which induces makeup flow of 2-3 |LlI7min.


A low-volume "T" with a glass window is designed to allow accurate alignment of the capillary in the junction. A 75-|im transfer line is attached to the other end of the T. A gap of 10-20 |Lim between the CE capillary and the transfer line was found to be ideal. Newer designs make this alignment procedure unnecessary (251). The makeup fluid is provided by filling a 1-mM syringe body with makeup fluid and attaching it to the top of the T (237).

The advantage of the liquid junction interface over the sheath-flow technique is that no syringe pump is required to provide the makeup volume. Conventional capillaries can be used as well. However, the sheath technique provided superior LODs and minimal band broadening (237). The band broadening is probably due to the presence of the junction itself, which has microliters of dead volume. The current was also higher using the liquid junction interface, and capillary washing was more difficult. The wash solution would exit the makeup line, the path of least resistance, rather than exiting the ion source.

4. Direct Electrode

This approach is similar to the sheathless technique in that no makeup flow is required. The capillary is inserted into two stainless steel capillaries. A gold wire is placed just inside the outlet side of the capillary, and it is attached to the outside steel capillary with some silver paint to make the electrical contact (109). Since specialty capillaries may be required, this approach may only find limited usage. What was significant about the cited work was the use of time-of-flight (TOE) mass spectrometry instead of the conventional quadropole.

C. FAST-ATOM BOMBARDMENT

East-atom bombardment (FAB) is a soft ionization method well suited for the determination of polar and thermally labile large molecules. Samples are introduced into the mass spectrometer, dissolved in glycerol, thioglycerol, or another nonvolatile material. This carrier is known as the EAB matrix. Once inside the high-vacuum region of the instrument, the sample is exposed to a beam of high-energy xenon atoms, causing molecules at the surface of the matrix to be ejected and ionized. High-resolution MS up to a mass of about 6000 Da is practical using this technique.

The sheath flow interface, shown in Figure 9.24, is best employed for introducing the FAB matrix mixed with the effluent from the separation capillary (92-94, 252-254). Flow rates of 1-5 jlL/min or less of FAB matrix are generally employed. Since the probe tip is inside the high-vacuum region of the spectrometer, about 50 pL/min of hydrodynamic flow is induced inside a 10-|Lim-i.d.


CZE Capillary Sheath Capillary

Stainless-Steel FAB Tip

Torr Seal

FLOWS

He Cooling Gas

CZE FAB Matrix

Vespel Stainless-Steel insulator Probe Shaft

Polypropylene Tubing

FIGURE 9.24 Schematic of a CZE coaxial FAB-MS tip. Reprinted with permission from J. Chro-matogr., 516, 167 (1990), copyright © 1990 Elsevier Science Publishers.

capillary (93). This does not contribute substantially to band broadening and can actually be used for sample injection.

Both positive- and negative-ion modes are operative, depending on the nature of the matrix media. For positive-ion formation (Figure 9.25), heptafluorobu-tyric acid, pH 3.5, can be used (242); for negative-ion generation, ammonium hydroxide is useful (255). These modifiers also serve to increase the conductivity of the FAB matrix, which is necessary to maintain electrical contact between the FAB probe tip and the CE capillary (255). Like electrospray, the probe tip also serves as the electrical ground of the system (93). The probe tip is maintained at ±8 kV, leaving ±22 kV for the electrophoretic separation. Negative-ion detection is about an order of magnitude less sensitive than the positive-ion mode. Separation as negative ions followed by detection as positive ions can help overcome this problem for zwitterionic peptides and proteins (255). Volatile buffers such as 5 mM ammonium acetate or 10 mM acetic acid are useful for CE-FAB. Phosphate buffers can decrease the total ion current (93). FAB does not tolerate high-salt-content buffers, and so treated capillaries should be used to minimize wall interactions. Alternatively the salts can be removed from the "detector-side" buffer and replaced with FAB matrix solution (92). This discontinuous buffer system appears to function satisfactorily.

The sensitivity of FAB can be as much as two to three orders of magnitude poorer than electrospray, especially for smaller molecules. For small peptides, the CLOD is about 10"^ M (92). The advantages of FAB over electrospray are: (i) easy interface to a high-resolution mass spectrometer; and (ii) freedom from multiple charge, resulting in a simplified mass spectrum. The production of high-resolution mass spectra from tandem experiments provides for more clear-cut structural assignments compared with those run on a quadrupole instru-

9.10 Micropreparative Fraction Collection 405

ment. Despite these advantages, it appears that electrospray has greater general utihty as a CE-MS interface.

9.10 MICROPREPARATIVE FRACTION COLLECTION

A. INTRODUCTION

Fraction collection by CE bears little resemblance to the analogous process employed in HPLC. With the electroosmotic flow rate seldom greater than 100 nL/min, droplets at the end of the capillary do not form, since the rate of evaporation is greater than the rate of droplet formation. Despite these problems, nearly 100 papers have appeared in this area.

The problem of nonuniform migration velocity also exists and is similar to that described in Section 9.1. Both the lag times from the detection point and

FIGURE 9.25 Single ion electropherograms of 2 x 10"^ M solutions of the protonated molecular ions of (A) morphiceptin (30 fmol, N = 91,000), (B) proctolin (24 fmol, N = 120,000), and (C) Phe-Leu-Glu-Glu-Ile (24 fmol, N = 120,000). Capillary: 15-^im-i.d., length not specified; buffer: 5 mM ammonium acetate, pH 8.5; FAB matrix: glycerohwater (25:75) adjusted to pH 3.5 with heptafluorobutyric acid; FAB matrix flow rate: 0.5 |LiL/min; injection: electrokinetic, 6 kV for 2 s. Reprinted with permission from J. Chromatogr., 516, 157 (1990), copyright © 1990 Elsevier Science Publishers.


the appropriate collection time of each fraction are directly related to migration velocity for each component.

In addition, one cannot ignore the minute quantities of material that are separated in a single run. Multiple runs are generally required in order to collect sufficient quantities of material, particularly if a trace component needs to be identified. The potential for electrochemical reactions at the surface of the electrode in the receiving electrolyte must be considered as well.

The first reports employing fraction collection appeared in 1985 (256), and this technique was further refined a few years later (257, 258). These papers described a system where the voltage was interrupted each time the capillary-electrode assembly was moved to a new receiver vial. Most commercial instruments use a variation of this procedure.

B. PERFORMING A RUN

For instruments containing both an autosampler and a "fraction collector," such as the Beckman P/ACE 2000 series, the Hewlett Packard HPCE^^, and the Bio-Rad BioFocus 3000, the following procedure applies:

1. Perform an analytical run to establish the migration times for each solute. Ensure that the migration times are reproducible. Adjust the separation conditions so that the peaks of interest are separated by at least 10-20 s.

2. Calculate the number of runs required to produce a sufficient mass of material. Refer to Eqs. (8.4) and (8.7) and Table 8.3 to make the appropriate calculations. When performing multiple runs, change the buffer solution regularly to minimize buffer depletion. As many as 50 runs may be necessary to collect a sufficient amount of a trace component. Changing the buffer every 10 runs is generally sufficient.

3. Calculate the time required for the leading and tailing edge of each fraction to traverse the length from the detector to the end of the capillary. First, determine the migration velocity v^, using

V^ = ^ . (9.9)

Then the lag time for the leading and tailing edge of each component, L, is given by

t, = ^ , (9.10)

where Lf = the length of the capillary from the detector to the fraction collector.

9.10 Micropreparative Fraction Collection 407

4. Fill the receiver containers with 5-50 |XL of buffer. Use the smallest volume vials possible. Both the capillary and electrode must be submerged in buffer solution. Electrochemical reactions become more likely when extremely small collection volumes are employed. When working with small collection volumes, cooling of the fraction collector may be required to lower the evaporation rate.

5. After the run, evaporate or lyophilize to reduce the fraction volume, if necessary.

C. OPTIMIZING THE AMOUNT OF MATERIAL COLLECTED

Several combinations and compromises are possible to maximize the mass of material collected with each run.

1. Use large-i.d. capillaries such as a 100-|im- (259), a 180-|Lim- (260), or a 200-|im-i.d. capillary (261). Reduce the field strength by following an Ohm's law plot to minimize heating effects. Beware of siphoning effects and buffer depletion when large-diameter capillaries are used (260).

2. Increase the buffer ionic strength. High-ionic-strength buffers permit the loading of more concentrated sample solutions without loss of resolution (see Chapter 10). To compensate for Joule heating, lower the capillary temperature to subambient if possible (259). Reduce the field strength as well.

3. Lower the EOF to improve resolution. Coated capillaries, linear alcohols, or the use of DjO as the buffer diluent (262) may be useful here without elevating the current.

4. Consider the use of CITP as the mode of separation. This technique stacks only dilute material and permits very large injections to be made (263).

5. The use of rectangular capillaries has been reported to increase the loading capacity by a factor of three (264).

6. For proteins and peptides, CIEF can be utilized (265). 7. When collecting peaks that are closely spaced, reducing the voltage near the

collection time can be used to widen the time window This technique has been used when collecting oligonucleotides from gel-filled capillaries (266).

D. INSTRUMENTS WITHOUT FRACTION COLLECTORS

A clever scheme developed by Albin et ah (267) permits the use of the autosam-pler as a fraction collector. Using this method, it is possible to collect 10-20 pmol peptide in 4 runs using a 75-|im-i.d. capillary or in 10 runs with a 50-|am-i.d.


capillary if the starting sample is 100 nmol/mL or greater (268). The procedure is illustrated in Figure 9.26.

1. Perform an analytical run to calculate migration times and migration time precisions of all components to be collected.

2. Inject a small plug of water, followed by sample and then another small water plug. Bracketing by water allows the sample to restack following hydrodynamic band broadening to the detector (step 3).

3. Apply vacuum to draw the sample plug just beyond the detector window. 4. Reverse the power supply polarity and perform the separation in the direc

tion toward the autosampler. Trigger the autosampler for fraction collection as determined by the individual migration times of each component. Collect in 5-10 |LiL buffer solution. Cool the autosampler to minimize evaporation.

CAPILLARY

FLUSH/EQUILIBRATE 20" VACUUM

AUTOSAMPLER

DETECTOR

INJECT WATER/SAMPLE/WATER 5V5V5" or 5VEK/5"

INJECTION

e PULL SAMPLE PLUG PAST DETECTOR 5" VACUUM

m PERFORM SEPARATION -

REVERSE POLARITY

13: 3E AUTOSAMPLER/ FRACTION

COLLECTOR

FIGURE 9.26 Illustration of the procedure for fraction collection in the autosampler of an automated CE instrument. Reprinted with permission from Applied Biosystems.

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223. Cai, J., Henion, J. Capillary Electrophoresis-Mass Spectrometry. J. Chromatogr, A, 1995; 703:667.

224. Garcia, E, Henion, J. Fast Capillary Electrophoresis-Ion Spray Mass Spectrometric Determination of Sulfonylureas. J. Chromatogr, 1992; 606:237.

225. Henion, J. D., Mordehai, A. V, Cai, J. Quantitative Capillary Electrophoresis-Ion Spray Mass Spectrometry on a Benchtop Ion Trap for the Determination of Isoquinoline Alkaloids. Anal. Chem., 1994; 66:2103.

226. Hseih, E Y. L., Cai, J., Henion, J. Determination of Trace Impurities of Peptides and Alkaloids by Capillary Electrophoresis-Ion Spray Mass Spectrometry. J. Chromatogr, A, 1994; 679:206.

227. Huang, E. C, Wachs, T., Conboy J. J., Henion, J. D. Atmospheric Pressure Ionization Mass Spectrometry: Detection for the Separation Sciences. Anal. Chem., 1990; 62:713.

228. Lamoree, M. H., Tjaden, U. R., van der Greef, J. Online Coupling of Micellar Electrokinetic Chromatography to Electrospray Mass Spectrometry. J. Chromatogr, A, 1995; 712:219.

229. Takada, Y., Sakairi, M., Koizumi, H. Online Combination of Micellar Electrokinetic Chromatography and Mass Spectrometry Using an Electrospray-Chemical Ionization Device. Rapid Commun. Mass Spectrom., 1995; 9:488.

230. Muijselaar, R G., Yokio, M., Otsuka, K., Terabe, S. Online Partial Tilling Micellar Electrokinetic Chromatography-Mass Spectrometry. Chromatography, 1997; 18:136.

231. Nelson, W M., Lee, C. S. Mechanistic Studies of Partial-Filling Micellar Electrokinetic Chromatography. Anal. Chem., 1996; 68:3265.

232. Yang, L., Harrata, A. K., Lee, C. S. On-Line Micellar Electrokinetic Chromatography-Elec-trospray Ionization Mass Spectrometry Using Anodically Migrating Micelles. Anal. Chem., 1997; 69:1820.

233. Yang, L., Lee, C. S. Micellar Electrokinetic Chromatography-Mass Spectrometry. J. Chromatogr, A, 1997; 780:207.

234. Tanaka, Y, Kishimoto, Y, Otsuka, K., Terabe, S. Strategy for Selecting Separation Solutions in Capillary Electrophoresis-Mass Spectrometry. J. Chromatogr, A, 1998; 817:49.

235. The API Book. 1990, Sciex. 236. Johansson, I. M., Huang, E. C, Henion, J. D., Zweigenbaum, J. Capillary Electrophoresis-Atmo-

spheric Pressure Ionization Mass Spectrometry for the Characterization of Peptides. Instrumental Considerations for Mass Spectrometric Detection. J. Chromatogr, 1991; 554:311.

237. Pleasance, S., Thibault, P, Kelly, J. Comparison of Liquid-Junction and Coaxial Interfaces for Capillary Electrophoresis-Mass Spectrometry with Application to Compounds of Concern to the Aquaculture Industry. J. Chromatogr, 1992; 591:325.

238. Ramsey, R. S., McLuckey S. A. Capillary Electrophoresis/Electrospray Ionization Ion Trap Mass Spectrometry Using a Sheathless Interface. J. Microcolumn Sep., 1995; 7:461.


239. Lazar, I. M., Xin, B., Lee, M. L., Lee, E. D. Design of a Time-of-Flight Mass Spectrometer as a Detector for Capillary Electrophoresis. Anal. Chem., 1997; 69:3205.

240. Smith, R. D., Loo, J. A., Edmonds, C. G., Barinaga, C. J., Udseth, H. R. New Developments in Biochemical Mass Spectrometry: Electrospray Ionization. Anal. Chem., 1990; 62:882.

241. Thibault, R, Paris, C , Pleasance, S. Analysis of Peptides and Proteins by Capillary Elec-trophoresis/Mass Spectrometry Using Acidic Buffers and Coated Capillaries. Rapid Commun. Mass Spectrom., 1991; 5:484.

242. Smith, R. D., Loo, J. A., Edmonds, C. G., Barinaga, C. J., Udseth, H. R. Sensitivity Considerations for Large Molecule Detection by Capillary Electrophoresis-Electrospray Ionization Mass Spectrometry. J. Chromatogr., 1990; 516:157.

243. Banks, J. E Recent Advances in Capillary Electrophoresis/Electrospray/Mass Spectrometry. Electrophoresis, 1997; 18:2255.

244. Smith, R. D., Udseth, H. R. Capillary Zone Electrophoresis-Mass Spectrometry Nature, 1988; 331:639.

245. Wheat, T. E., Lilley K. A., Banks, J. E Capillary Electrophoresis with Electrospray Mass Spectrometry Detection for Low-Molecular-Mass Compounds. J. Chromatogr, A, 1997; 781:99.

246. Varesio, E., Cherkaoui, S., Veuthey, J.-L. Optimization of CE-ESI-MS Parameters for the Analysis of Ecstasy and Derivatives in Urine. J. High Resolut. Chromatogr, 1998; 21:653.

247. Alexander, J. N., Schultz, G. A., Polil, J. B. Development of a Nano-electrospray Mass Spectrometry Source for Nanoscale Liquid Chromatography and Sheathless Capillary Electrophoresis. Rapid Commun. Mass Spectrom., 1998; 12:1187.

248. Cao, P, Moini, M. Analysis of Peptides, Proteins, Protein Digests, and Whole Human Blood by Capillary Electrophoresis/Electrospray lonization-Mass Spectrometry Using an In-Capillary Electrode Sheathless Interface. J. Am. Soc. Mass Spectrom., 1998; 9:1081.

249. Mazereeuw, M., Hofte, A. J. P., Tjaden, U. R., van der Greef, J. A Novel Sheathless and Elec-trodeless Microelectrospray Interface for the Online Coupling of Capillary Zone Electrophoresis to Mass Spectrometry. Rapid Commun. Mass Spectrom., 1997; 11:981.

250. Lee, E. D., Muck, W, Henion, J. D., Covey, T. R. On-Line Capillary Zone Electrophoresis-lon Spray Tandem Mass Spectrometry for the Determination of Dynorphins. J. Chromatogr, 1988; 458:313.

251. Wachs, T., Sheppard, R. L., Henion, J. Design and Applications of a Self-Aligning Liquid Junc-tion-Electrospray Interface for Capillary Electrophoresis-Mass Spectrometry. J. Chromatogr, B: Biomed. Appl, 1996; 685:335.

252. Caprioli, R. M. Continuous-Flow East Atom Bombardment Mass Spectrometry. Anal. Chem., 1990; 62:477.

253. Moseley M. A., Deterding, L. J., Tomer, K. B., Jorgenson, J. W. Capillary Zone Electrophoresis-Mass Spectrometry Using a Coaxial Continuous-Flow Fast Atom Bombardment Interface. J. Chromatogr, 1990; 516:167.

254. Wolf, S. M., Vouros, P., Norwood, C , Jackim, E. Identification of Deoxynucleotiside-Poly-aromatic Hydrocarbon Adducts by Capillary Electrophoresis-Continuous Flow-Fast Atom Bombardment Mass Spectrometry. J. Am. Soc. Mass Spectrom., 1993; 3:757.

255. Moseley, M. A., Deterding, L. J., Tomer, K. B., Jorgenson, J. W Determination of Bioactive Peptides Using Capillary Zone Electrophoresis/Mass Spectrometry Anal. Chem., 1991; 63:109.

256. Hjerten, S., Zhu, M.-D. Micropreparative Version of High-Performance Electrophoresis. The Electrophoretic Counterpart of Narrow-Bore High-Performance Liquid Chromatography. J. Chromatogr, 1985; 327:157.

257. Rose, D. J., Jorgenson, J. W. Fraction Collector for Capillary Zone Electrophoresis. J. Chromatogr, 1988; 438:23.

258. Cohen, A. S., Najarian, D. R., Paulus, A., Guttman, A., Smith, J. A., Karger, B. L. Rapid Separation and Purification of Oligonucleotides by High-Performance Capillary Gel Electrophoresis. Proc. Natl. Acad. Scl, 1988; 85:9660.

References 421

259. Grimm, R., Herold, M. Micropreparative Single Run Fraction Collection of Peptides Separated by CZE for Protein Sequencing. J. Capillary Electrophor., 1994; 1:79.

260. Yin, H., Keely-Templin, C, McManigill, D. Preparative Capillary Electrophoresis with Wide-Bore Capillaries. J. Chromatogr., A, 1996; 744:45.

261. Chen, E A., Kelly L., Palmieri, R., Biehler, R., Schwartz, H. Use of High Ionic Strength Buffers for the Separation of Proteins and Peptides with Capillary Electrophoresis. J. Liq. Chromatogr., 1992; 15:1143.

262. Camilleri, P., Okafo, G. N., Sou than, C, Brown, R. Analytical and Micropreparative Capillary Electrophoresis of the Peptides from Calcitonin. Anal. Biochem., 1991; 198:36.

263. Schwer, C , Lottspeich, E Analytical and Micropreparative Separation of Peptides by Capillary Zone Electrophoresis Using Discontinuous Buffer Systems. J. Chromatogr, 1992; 623:345.

264. Cifuentes, A., Xu, X., Kok, W. T., Poppe, H. Optimum Conditions for Preparative Operation of Capillary Zone Electrophoresis. J. Chromatogr, A, 1995; 716:141.

265. Foret, E, Muller, O., Thome, J., Gotzinger, W, Karger, B. L. Analysis of Protein Fractions by Micropreparative Capillary Isoelectric Focusing and Matrix-Assisted Laser Desorption Time-of-Flight Mass Spectrometry J. Chromatogr, A, 1995; 716:157.

266. Guttman, A., Cohen, A. S., Heiger, D. N., Karger, B. L. Analytical and Micropreparative Ultrahigh Resolution of Ohgonucleotides by Polyacrylamide Gel High-Performance Capillary Electrophoresis. Anal. Chem., 1990; 62:137.

267. Albin, M., Chen, S. M., Louie, A., Pairaud, C, Colbum, J. The Use of Capillary Electrophoresis in a Micropreparative Mode: Methods and Apphcations. Anal. Biochem., 1992; 206:382.

268. Hathaway, G. M. Preparative Capillary Electrophoresis for Offline Sequence, Composition, and Mass Analysis of Peptides. Methods, 1992; 4:244.

CHAPTER l O Putting It All Together

10.1 Selecting the Mode of HPCE 10.2 Requirements for Robust Separations 10.3 Realistic Compromises 10.4 Quantitative Analysis 10.5 Sample Preparation 10.6 Mobility as a Qualitative Tool 10.7 Validation 10.8 Troubleshooting

References

10.1 SELECTING THE MODE OF HPCE

Selecting the appropriate separation mode is straightforward for most applications. Table 10.1 contains some guidelines to suggest which technique will yield adequate results in the shortest period. The techniques are listed in descending order from the most to least favored. The basis for these assignments is ease of methods development, robustness of the separation process, and the appropriateness of the mode of operation to the problem to be solved.

Rugged and precise methods are generally simple in their design. For CZE separations, start by using a buffer such as borate or phosphate. Based on

Table 10.1 Selecting the Mode of Capillary Electrophoresis^

Small Ions

CZE

MECC

Small Molecules

CZE

MECC

Peptides

CZE

MECC

CIEF

PN

Proteins

CZE

CIEF

PN

MECC

Oligonucleotides

MECC

PN

CGE

DNA

PN

^CITP is not considered here. PN = polymer network

423

4 2 4 Chapter 10 Putting It All Together

experimental observations, the appropriate additives and coatings can then be used to properly design the separation system. For a well-controlled separation, expect run-to-run migration time RSDs as low as 0.5% and externally standardized peak area RSDs as low as 1%. Through the use of an EOF marker and an internal standard, both mobility and peak area precision improve yet further.

10.2 REQUIREMENTS FOR ROBUST SEPARATIONS

Poor reproducibility of separations has been a frequent problem in HPCE, particularly among new users. The following guidelines can serve as a checklist that is applicable to all methods. The troubleshooting guide in Section 10.8 fills in the details.

1. Temperature control The capillary must be thermostated. Most commercial instruments perform this task quite well. In some cases, the outlet side is not thermostated at all. Since viscosity and, thus, mobility are temperature dependent, failure to adequately control the capillary temperature will result in poor migration time precision.

2. Solubility. All sample components must be soluble in the BGE. This is particularly important whenever the injection solution composition differs from the supporting electrolyte. Pay careful attention to the buffer. Some solutes are not very soluble in phosphate buffer. In these cases, acetate buffer, pH 4, may be a better choice than phosphate buffer, pH 2.5. Try to avoid using organic solvents.

3. Wall effects. Capillary coatings or buffer additives may be required to suppress wall interactions. If solutes are adhering to the capillary wall, this problem must be dealt with prior to undertaking any further optimization.

4. Joule heating. Perform an Ohm's law plot to optimize the voltage. 5. Selectivity. The appropriate mobility plots should be performed to

select the optimal buffer composition for separation. Variables include pH and additive concentration. Rugged separations are generally obtained when small changes in buffer composition do not dramatically affect mobility. This information is readily discernible from mobility plots.

6. Buffer refreshment. Buffer depletion can result in drift of both mobility and EOF The sensitivity of this effect is solute dependent and may be based on the factors discussed above. The requisite frequency for buffer replenishment should be determined experimentally. Migration time drift is frequently observed from this effect, and resolution can be affected as well.

7. Evaporation control. Use closures provided by the instrument manufacturers for samples and buffers. Reduce the sampler temperature if your

10.4 Quantitative Analysis 425

instrument has that capabihty. Evaporation can be particularly troublesome when organic solvents are employed.

8. Control of the ionic strength of the injection solution. The conductivity of the injection solution including the sample contribution should be equal to or less than the conductivity of the BGE. Precise intersample control of the ionic strength is required. Beware of sample matrix effects. It may be necessary to perform a sample cleanup to place the solutes in an HPCE-friendly (e.g., low-ionic-strength) matrix. This effect becomes more important as the injection size is increased.

9. Sample viscosity control. The sample viscosity can affect the amount of material introduced into the capillary. Dilution or sample cleanup may be required to ensure that all samples have the same viscosity

10. Internal standards. An internal standard can compensate in part for quantitative problems that occur because of sample evaporation, injection variance, or sample-to-sample viscosity differences. The internal standard should be used to adjust for minor experimental variations. If substantial migration time or peak area variations occur, these should be corrected for experimentally.

11. Capillary washing. To ensure a clean and reproducible inner wall, it is often necessary to wash the capillary between runs with 0.1 N or even 1 N sodium hydroxide. A 1- or 2-min wash is usually adequate, followed by a 1-min water rinse and 1- to 2-min reequilibration with run buffer. This is only recommended for bare silica capillaries. For low-pH, low-EOF separations, a 2-min 0.1 N phosphoric or hydrochloric acid wash helps to maintain a low and reproducible EOE

12. Capillary inlet. Ensure that the capillary inlet is cut squarely and no poly-imide shards protrude off the end.

10.3 REALISTIC COMPROMISES

Speed, resolution, and loadability often oppose each other in separations. The acute sensitivity problem in HPCE must be factored in as well. As a starting point, a reasonable separation must have already been developed following the guidelines given in the individual chapters. Use Table 10.2 as a guide for optimizing a particular aspect of a separation.

Do not push these techniques too far. When heroic efforts are called for, the separation scheme is probably not appropriate to begin with.

10.4 QUANTITATIVE ANALYSIS

Examining of the early HPCE literature, it is easy to conclude that CE is a non-quantitative technique. This is particularly true prior to the advent of commercial

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instrumentation. Now that the dark ages of HPCE have passed, the technique is recognized as quantitative.

With fully automated instrumentation, online detection, and modem computing power for peak integration, quantitative analysis by HPCE is much simpler than for slab-gel electrophoresis. The precision and accuracy of HPCE far exceed those of the slab gel, and they are beginning to approach those of HPLC. In this section, the important factors governing quantitative analysis will be covered.

A. DATA SAMPLING RATE

Most data systems or integrators are suitable for HPCE, provided their data acquisition rates can be adjusted to account for the sharp peaks characteristic of HPCE. Sampling rates of 20 Hz or less are suitable for all but the most efficient separations. At that sampling rate, a signal with a 5-s peak width is sampled with 100 points, an amount sufficient for all necessary computations. Data collection of 25 points per peak is generally adequate. Oversampling does not substantially improve precision, but it does occupy disk storage capacity Under-sampling will dramatically increase the %RSD for peak area.

If there are any doubts about the integrity of the data system, it should be validated using an electronic peak generator. The error contribution of the data system should be negligible. Also be sure that the integration parameters are properly adjusted. Note the placement of start/stop integration tick marks, and be sure each peak is properly integrated. Poor resolution greatly increases integration error, since it is more difficult for the integrator to accurately integrate the peaks and properly identify the baseline.

B. USE OF PEAK AREAS

Peak areas, rather than peak heights, are best used in quantitative analysis for two reasons:

1. Variation in the ionic strength of the sample can result in more or less solute stacking. Both peak height and peak width are greatly affected. As long as resolution remains adequate, peak area remains constant.

2. The linear dynamic range of the separation is enhanced. As the solute concentration increases, electromigration dispersion begins to appear. Again, peak height and width are affected, but the area is conserved. This will be discussed in more detail later in the chapter.

When the sample is carefully controlled and the linear dynamic range is narrow, peak heights are reproducible.


C. EXTERNAL STANDARDS

If external standards are used for quantitative analysis, several sample parameters must be carefully controlled. These include sample viscosity, sample ionic strength, and sample temperature. If these factors vary from sample to sample, severe quantitative errors will occur. If samples were stored in the refrigerator overnight, it is important to allow them to come to room temperature prior to separation.

When following the aforementioned guidelines, external standardization works well. For example, in the determination of nonsteroidal anti-inflammatory drugs, the sample preparation involved sonicating an enteric-coated tablet for 10 min in methanol (1). The methanolic solution was diluted with BGE to yield a final drug concentration of 100 |Xg/mL. Since the amount of drug per tablet was a high as 500 mg, the methanohc solution was 5 mg/mL. A 50-fold dilution was then performed using BGE. In this case, it is fairly certain that each prepared sample will have the same viscosity. The results ranged from 101.5% to 104% recovery of the labeled amount of drug.

External standard calculations are performed by conventional methods, identical to those used in HPLC or GC. Calibration can be single point, but multipoint regression is usually preferred. For single-point calibration, the standard should have a solute concentration higher than that of any sample.

Area corrections for migration time scatter are generally not used. When the migration times vary by less than 1%, these corrections do not improve analytical precision. Data that support these conclusions can be found in Table 10.3a. In that study, the same separation was run in seven laboratories located in the United Kingdom. The migration time %RSDs ranged from 0.2% to 1.3%, with six of seven laboratories reporting better that 1% RSD (Table 10.3b). The peak area %RSDs ranged from 0.6% to 2.6%. When the peak areas were normalized by dividing by the migration time, no clear trend emerged. In some cases, precision improved; in others, though, there was no change or precision worsened. In the single case where the migration time %RSD was greater than 1%, the correction appeared useful.

Since this data set was small, larger scale studies might discover subtle trends. If the migration times fluctuate wildly area correction will probably help, but the analyst is far better off if the migration time precision is brought under control.

For single-point calibration, the concentration of a solute can simply be determined by

(A ^ r — SAMP ^SAMP ~

QTH , (10.1)

where CSAMP i d C STD ^re the respective concentrations of sample and standard solutions, and ASAMP i d ^STD ^ ^ the respective peak areas.


Table 10.3A Intercompany Cross-Validation Study for the Chiral Recognition of Clenbuterol: Peak Area Precision (n = 10)

Company

1

2

3

4

5

6

7

Precision, Peak 1 (%RSD)

Actual

1.2

2.6

0.6

1.3

2.2

2.5

1.7

Normalized

0.8

2.5

1.0

1.2

2.1

1.1

1.7

Precision, Peak 2 (%RSD)

Actual

1.4

1.9

0.4

1.7

2.2

2.9

1.5

Normalized

0.8

1.8

1.1

1.6

2.0

1.8

1.5

Peak Area Ratio

%RSD

0.2

0.5

0.4

0.9

0.3

0.6

0.9

Experimental Conditions Capillary: 57 cm x 50 pm i.d.; BGE: 30 mM hydroxypropyl-j3-cyclodextrin in 50 mM borate adjusted to pH 2.2 with cone, o-phosphoric acid; injection: 8 nL; voltage: 30 kV; detection: UV, 214 nm; temperature: ambient; intersample wash: 1 min with 0.1 N sodium hydroxide; BGE: reequilibration: 2 min; solute: clenbuterol racemate, 150 pg/mL in water. Data is reproduced from reference (88),

For multiple-point calibration, linear regression is used to determine the best line through the calibration data. For nonlinear response, quadratic or polynomial curve fitting can be employed, but this is less dependable than a linear fit. Standards should bracket the expected solute concentration range. Linearity will be discussed in more detail shortly.

Table 10.3B Intercompany Cross-Validation Study for the Chiral Recognition of Clenbuterol: Migration Time Precision (n = 10)

Company Instrument Migration Time (%RSD)

Beckman

Beckman

Beckman

Spectra-Physics

ABl

Beckman

ABI

1.3

0.3

0.8

0.4

0.6

0.5

0.2


Table 10.3C Intercompany Cross-Validation Study for the Chiral Recognition of Clenbuterol: Peak Area Linearity, 6 Levels, 10-150%

Company Correlation Coefficient y Intercept

1 0.999 0.98

2 0.997 0.31

3 0.999 -0.33

4 0.992 5.10

5 0.996 0.33

6 0.990 -1.34

7 0.994 3.53

D. INTERNAL STANDARDS

Internal standards are added to the sample to correct for quantitative losses during cleanup as well as instrumental imprecision, primarily due to the injection process. Calibration can be single or multilevel, with the latter the preferred mode. Using a standard solution, the ratio (R) of peak areas of the standard and internal standard is calculated. If the same amount of internal standard is added to a sample, then

(10.2)

where CSAMP ^d CSJD are the respective concentrations of sample and standard solutions, and RSAMP and R^j^ are the respective peak area ratios.

The internal standard must be used to compensate for sample cleanup losses if quantitative recoveries cannot be made. More importantly, the internal standard compensates for differences in sample viscosity, in temperature, and in the instrumental error from injection. This is illustrated using data shown in Table 10.3a. Peak 1 and peak 2 represent the optical isomers of clenbuterol. If peak 2 is considered the internal standard, when the %RSD of the peak area ratio is calculated, the range is from 0.2% to 0.9%. This clearly indicates the value of the internal standard.

For quality control purposes, the internal standard is invaluable. If a peak appears at the correct migration time with the correct peak area, it indicates that the instrument is functioning appropriately. The use of a neutral marker is also helpful in this regard when the EOF is high. In this case, the neutral marker


indicates any changes in the EOF. When the EOF is low, the neutral marker is less useful, since its measurement will usually prolong the time of separation.

For the modes of CIEF and CGE, internal standards should be added to correct for migration time variations, due in part to the salt content of the sample. This same internal standard can be used for quantitation as well.

E. AREA PERCENT CALCULATIONS

Area percentage is frequently used for the purity determination of bulk pharmaceuticals. The areas of all peaks are summed, and the percent contribution of each component is calculated. The method is based on the assumption that potential process impurities and degradation products have identical molar absorbtivities at the wavelength of detection, usually in the low UV For this application, it is critical to correct the peak areas of the parent compound and its impurities for their migration velocities. When uncorrected areas are used, positive or negative errors will occur, depending the relative migration order of the sample's components.

E LINEAR DYNAMIC RANGE

Nonlinear effects can be observed in HPCE under several conditions. With elec-trokinetic injection, as the solute concentration is increased, the ionic strength increases as well. This reduces the field strength over the sample, and as a result, a negative deviation from linearity will occur. The problem of electrodispersion can also be observed. This will affect the linear dynamic range for either hydro-dynamic or electrokinetic injection.

The results obtained for the serial dilution of naproxen starting at 10 mg/mL are shown in Figure 10.1 (1). At that high concentration, a badly shaped broadened peak is obtained. As the serial dilution proceeds, the peak width narrows from approximately 24 s to 3 s. The cause of the peak broadening is electrodispersion (Section 2.14). Once the solute concentration is below 156 |Xg/mL, a consistent peak is obtained. Note that the point of inflection of the peak shifts to longer times as the solute concentration is increased. The migration time reproducibility was incredibly good that day, as evidenced by the three runs at the lowest concentrations. It is probable that the increase in migration time from 2.55 min to 2.56 min is significant and represents the onset of electrodispersion.

The figures of merit from these runs are plotted in Figure 10.2. Above 100 |lg/mL, deviations from expected values for migration time, peak width,

432 Chapter 10 Putting It All Together

CONC.(ug/mL)

156 -

78

0.078 9 . 8

FIGURE 10.1 Impact of solute concentration on peak shape. Capillary: 60 cm (38 cm to detector) X 50 |Lim i.d.; buffer: 25 mM SDS, 20 mM borate, pH 9.2; temperature: 50°C; voltage: 25 kV; detection: UV, 230 nm; injection: vacuum, 1 s; solute: naproxen. Reprinted with permission from J. Liq. Chromatogr., 14, 953 (1991), copyright © 1991 Marcel Dekker.

and peak height are evident. Peak area shows less deviation from linearity. This is quite reasonable, since at high solute concentrations, much of the signal is represented in peak width rather than in peak height. To maximize the linear dynamic range of the separation, it is important to use peak areas for


10

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mg/ml

FIGURE 10.2 Impact of solute concentration on analytical figures of merit. Refer to Figure 10.1 for experimental conditions. Key: Log A = log peak area; Log HGT = log peak height; (MX -2) X 10~ = scaled migration time (min); PW/3 = scaled peak width (s). Reprinted with permission from J. Liq. Chromatogr., 14, 953 (1991), copyright © 1991 Marcel Dekker.

all calculations. Results from an interlaboratory cross-validation study are given in Table 10.3c. The data indicate that linearity is reproducible in multiple laboratories.

Increasing the linear range is possible with high-ionic-strength buffers (1). Figure 10.3 shows separations of some anti-inflammatory drugs at concentrations of 1 mg/mL and 250 |ig/mL using a low-ionic-strength buffer. Substantial losses in resolution are found at the higher solute concentrations. A similar separation in a high-ionic-strength buffer is shown in Figure 10.4. Almost no change in resolution is found between the run with high concentration and that with low concentration. Note that a shortened capillary was employed, since the EOF was substantially lowered by the high-ionic-strength buffer. The separations in these two figures were performed in 25-|im-i.d. capillaries to reduce the heating effects.

The overall addressable concentration range of HPCE is illustrated in Figure 10.5 on p. 436. The use of the laser detector solves most of the compelling problems in HPCE. With this highly sensitive detector, it is possible to perform extreme dilutions of most samples. At high dilution, most ionic-strength-mediated effects from the solute and/or the sample matrix become insignificant. Unfortunately, derivatization is required for most laser-fluorescence-based applications.


1.6

FIGURE 10.3 Band profile dependence on solute concentration and buffer ionic strength for low-ionic-strength buffer. Capillary: 65 cm (43 cm to detector) x 25 im i.d.; buffer: 20 mM SDS, 20 mM phosphate, pH 9.2; temperature: 30°C; injection: vacuum, 2 s; detection: UV, 230 nm. Key: (A) (1) suhndac, 1 mg/mL; (2) indomethacin, 1 mg/mL; (3) tolmetin, 1 mg/mL; (4) ibupro-fen, 1 mg/mL; (5) naproxen, 100 |Llg/mL; (6) diflunisal, 500 |lg/mL. (B) 4x dilution of A. Reprinted with permission from J. Liq. Chromatogr., 14, 953 (1991), copyright © 1991 Marcel Dekker.

10.5 SAMPLE PREPARATION

A. BASIC PRINCIPLES

In HPCE, there are several important issues regarding sample preparation. Obviously, interfering components, if not separable, must be removed during the sample preparation process. This is true for all analytical techniques. In both chromatography and electrophoresis, the sample matrix can affect the resolution

10.5 Sample Preparation 435

L

FIGURE 10.4 Band profile dependence on solute concentration and buffer ionic strength for high-ionic-strength buffer. Conditions as per Figure 10.3 except: capillary: 20 cm to detector; buffer: 100 mM phosphate, 25 mM SDS, pH 7.0. Reprinted in part with permission from J. Liq. Chromatogr., 14, 953 (1991), copyright © 1991 Marcel Dekker.


LASER NORMAL MICRO STATE-OF FLUORESCENCE STACKING ANALYTIC PREPARATIVE THE-ART DETECTION RANGE RANGE RANGE

10""' 10" ^ 10"''° 10" 10" 10" 10' CONCENTRATION (M)

FIGURE 10.5 The dynamic ranges of capillary electrophoretic techniques.

of the separation. Unlike chromatographic techniques, in HPCE the sample matrix may have a profound impact on the amount of material that is injected into the capillary when electrokinetic injection is employed. The sample preparation process must deal with this problem.

For samples containing high concentrations of solutes—for example, pharmaceutical dosage forms—simple dilution of the dosage form extract in the supporting electrolyte is sufficient, since generally only small injection volumes are required. Depending on the strength of the chromophore, final concentrations of 10"^-10"^ M provide adequate sensitivity.

For complex samples or when high sensitivity is required, sample preparation to remove interferences and place the solute(s) in a CE-friendly solution is clearly indicated. Stacking techniques can be useful for improving sensitivity, but matrix effects and artifacts often interfere. There are hundreds of examples in the literature describing these techniques, some of which will be cited in the following discussion. Centrifugation or filtration to remove particulate matter is always good practice. Sonication to remove air is sometimes needed as well.

In reversed-phase LC, it is generally bad practice to prepare the sample in a solvent with greater eluting power than the mobile phase, particularly if large injections are required. The fundamental requirement for HPCE is that the sample should never be prepared so as to have an ionic strength greater than that of the supporting electrolyte. This requirement can be loosened only when the injection volume can be kept small. It is usually good practice to desalt high-ionic-strength samples.

For small molecules, the most useful forms of sample preparation include:

1. Liquid-liquid extraction (2-9) 2. Solid-phase extraction (9-18) 3. Supercritical fluid extraction (19) 4. Protein precipitation (for blood serum or plasma) (20-26) 5. Dialysisi (27-29) 6. Ultrafiltration (30)

Un yivo microdialysis is also employed for sampling of neurochemicals. This subject is beyond the scope of this text.


For large molecules such as proteins and DNA, the following techniques are also applicable:

1. HPLC (31) 2. Affinity LC (32) 3. Ultrafiltration (33, 34) 4. Solid-phase extraction (35-41) 5. Dialysis (42, 43) 6. Desalting (44-47) 7. Sedimentation (48) 8. Precipitation (49-54)

In the following sections, some of these modes of sample preparation are covered in greater detail.

B. DRUGS IN BIOLOGICAL FLUIDS

1. Direct Injection

MECC is often preferred for separating small synthetic pharmaceuticals. Since surfactant solutions are utilized, direct injection of blood plasma or serum might be feasible, since a surfactant such as SDS binds strongly to and solubilizes serum proteins. Indeed, micellar liquid chromatography has been shown useful for direct injection. In this mode of HPLC, the surfactant solution serves as the mobile-phase modifier. Surfactant-bound serum proteins form an extremely large aggregate, which is excluded from the stationary phase. The protein bolus elutes on or about t^ in a relatively narrow band. The retained drug substance elutes some time later, producing clean chromatograms at the low micrograms-per-milliliter level.

In MECC, the protein-surfactant aggregate has a substantial net negative charge when SDS is used as the additive. The aggregate then elutes relatively late in the separation, leaving only a small window for interference-free monitoring of the drug substance. The advantage of MECC with direct injection over CZE with acetonitrile protein precipitation has been studied (55). The technique gave better interday precision (1.49% vs. 16.1%) than did CZE.

Direct injection will become successful only under one of certain circumstances:

1. Selective detection is possible. 2. The drug substance is present in serum or plasma at high concentrations.

Selective detection includes fluorescence and UV detection at wavelengths above 240 nm. When low UV detection is used, a CLOD of 5 jlg/mL is found (56). With liquid-liquid extraction, the CLOD drops to 1 |Llg/mL, and with solid-phase extraction, a CLOD of 100 ng/mL is obtained. The baselines are always cleaner when some form of sample preparation is employed.


Since low-UV detection is often required in HPCE, a typical separation is shown in Figure 10.6 for aspoxicillin at a concentration of 50 |Llg/mL (57). The problems of unambiguously assigning peak identity are immediately obvious. Nevertheless, direct injection is particularly useful when the sample size is extremely limited—for example, clinical determination of xanthines from premature infants (58).

y iw^

1 — 0

—r-5

1 10

TIME (min.)

1

15

— r 20

FIGURE 10.6 Direct plasma injection for the determination of aspoxicillin. Capillary: 65 cm (50 cm to detector) x 50 p,m i.d.; buffer: 50 mM SDS, 20 mM phosphate-borate, pH 8.5; voltage: 20 kV; detection: Uy 210 nm; temperature: ambient; solute concentration: 50 |J,g/mL. Reprinted with permission from J. Chromatogr., 515, 245 (1990), copyright © Elsevier Science Publishers.


2. Protein Precipitation

In the simplest form of the procedure, proteins can be precipitated by adding 100 |iL of acetonitrile to 200 |LIL of blood plasma or serum that already contains an internal standard. The mixture is then vortex mixed for 30 s, allowed to stand for 5 min at room temperature, and centrifuged for 3 min at 9500g, and the supernatant is injected (56). The electropherograms are cleaner than for direct injection, but many endogenous components still appear. MECC is still advantageous, since some proteinaceous material may carry over into the supernatant.

In conjunction with added salts to the sample, efficient stacking (Section 8.6) can be obtained (59-62). This is illustrated in Figure 10.7 (60). With the drugs dissolved in 67% acetonitrile-150 mM sodium chloride and 50% of the capillary filled with sample, efficient stacking is obtained (Figure 10.7a). The serum blank (Figure 10.7b) is relatively clean. The spiked serum sample is shown in Figure 10.7c. It is likely that tITP is the stacking mechanism at work and that chloride is the leading ion.

3. Liquid-Liquid Extraction

Liquid-liquid extraction is useful for performing an offline trace enrichment. This is illustrated for the determination of thiopental in serum and plasma (6). Buffered serum (0.7 mL) containing an internal standard was extracted with 5 mL of pentane for 10 min and centrifuged. The upper organic layer was removed and evaporated to dryness. The residue was redis-solved in 200 |lL of BGE, with separation by MECC. The electropherograms were free of endogenous sample peaks, and the results from patient samples correlated well with the HPLC assay. Should further enrichment and sensitivity be required, the pickup solvent volume can be reduced, and instead of using the BGE, a stacking electrolyte can be used.

4. Solid-Phase Extraction

Solid-phase extraction is a widely used sample-preparation method for purifying drugs from biological fluids prior to HPLC. Wernly and Thormann (14) employ multistep solid-phase extraction to determine drugs of abuse such as barbiturates, hypnotics, amphetamines, opiods, benzodiazepines, and cocaine metabolites from a single urine specimen. In conjunction with multiwavelength detection, positive confirmation for drugs of abuse in screening urine samples is simple by MECC. The stepwise sample cleanup procedure is illustrated in Figure 10.8. In this three-step approach, methaqualone is eluted during the first step, morphine, codeine, and heroin during the second, and finally, benzoylecgonine in the third. There was some carryover between fractions that should be readily eliminated through fine-tuning. It is likely that this procedure can be easily adapted for the determination of a wide variety of drug substances in most biological fluid types.

2

a

! ' 1 ' »i< I I I JU I I . I i I

-f r

i!

r^ 1 UUvJ.,.

Min

FIGURE 10.7 Electropherograms of (1) iohexol, (2) theophylline, and (3) phenobarbital. Capillary: 42 cm X 50 |Lim; BGE: 250 mM boric acid, pH 8.9; injection: 50% of capillary filled with sample; detection: UV, 214 nm; temperature: 30°C; field strength: 280 V/cm; sample preparation: 100 |xL of serum vortex mixed with 200 |iL of acetonitrile and centrifuged at 14,000g for 30 s, all solutions contained 150 mM sodium chloride; solute concentration: 10 |Llg/mL; (a) standards; (b) serum blank; (c) spiked serum. Reprinted with permission from J. Capillary Electrophor., 2, 267 (1995), copyright © 1995 International Scientific Communications.

10.5 Sample Preparation

Condition Sample application

Wash Elutlon A Vi sh

441

Eiution B Elutlon C

0^m

6 6 W^ste Waste

i i i i i

Waste Analysis

m

6 Waste

i ^p i M H

(s) 4 Analysis Analysis

Bonded phase packing M - Matrix compound A, B, C - Sampte compounds

FIGURE 10.8 Solid-phase extraction process for the determination of drugs in urine samples. (1) Condition with 2 mL each of methanol and 100 mM phosphate buffer, pH 6, just before use. (2) Load 5 mL urine-2 mL phosphate buffer via vacuum over a 2-min span. (3) Wash sequentially and discard solutions of 1 mL of phosphate buffer:methanol (80:20), 1 mL of 1 M acetic acid, and 1 mL of hexane. (4) Elute with 4 mL of methylene chloride. (5) Wash with 6 mL of methanol. (6) Elute with 2 mL of 2% ammonium hydroxide in ethyl acetate. (7) Elute with 2 mL of methylene chloride:isopropyl alcohol (80:20) containing 2-10% ammonium hydroxide. Reprinted with permission from And. Chem., 64. 2155 (1992), copyright © Am. Chem. Soc.

A detailed procedure for determining acidic drugs is as follows (9):

1. Condition the SPE cartridge with 10 mL of methanol followed by 10 mL of water at a flow rate of about 1 drop per second.

2. Mix 0.5 mL of whole blood containing an internal standard with 1 mL of 0.1 N HCl, and vortex for 15 s.

3. Transfer to a 3-mL syringe attached to the cartridge. 4. Flush the cartridge with 10 mM water. 5. Elute acidic drugs with 4 mL of ethyl acetate. 6. Centrifuge the ethyl acetate at 2000 rpm for 10 min. 7. Evaporate to dryness. 8. Redissolve the residue in 1:9 ethanoLwater. 9. Centrifuge to gather liquid at the bottom of the vial.

10. Transfer the material to an injection vial, cap, and centrifuge at 12,000 rpm for 10 min.

11. Cut off the vial cap, place the vial in a spring-loaded sample vial (Beck-man), and inject.


The advantages of these approaches are as follows:

1. Large sample sizes can be processed and trace enriched, solving, in part, some of the sensitivity problems.

2. The organic solvent eluting reagents are easily evaporated, after which the residue can be redissolved in a small quantity of run buffer or a stacking buffer if necessary.

3. The process produces relatively clean electropherograms.

C. PROTEINS AND D N A

The appropriate sample preparation for proteins or DNA depends on the sample matrix and the need for concentration of the sample. In many cases, no sample preparation is required other than filtration or centrifugation. When a chromatographic step or solid-phase extraction is used for purification of the sample, it is often necessary to desalt the solution prior to injection. Solid-phase extraction can also be used to desalt oligonucleotide samples (46). When the solute concentration is low, it may be necessary to concentrate the solute. Ultrafiltration or precipitation techniques provide both desalting and enrichment. Dialysis is used for desalting, purification, or buffer exchange.

1. Ethanol Precipitation of DNA

Ethanol precipitation of PCR-produced DNA is often used as a final purification step. The following is a typical protocol starting with DNA that has already been purified by agarose gel electrophoresis (53):

1. Mix 1 volume of purified DNA with 3 volumes of 70% ethanol. 2. Store for 2 h at-70°C. 3. Centrifuge at 20,000 rpm for 20 min at 4°C. 4. Remove supernatant, and dry pellet. 5. Redissolve in 10 |LiL of 1 mM Tris-HCl-0.1 mM EDTA. 6. Combine 2 |iL of sample with 3 |LiL of formamide and 5 |LiL of water. 7. Boil for 3 min to denature. 8. Quick-freeze in dry ice-acetone and store at -20°C until ready for

analysis.

2. Acetone Precipitation of Proteins

The following protocol is employed to concentrate and desalt a lavage fluid obtained from the lungs of rats exposed to perfluoroisobutylen (54). The proteins were swept from the lung by lavage with 150 mM sodium chloride.


1. Add 10 volumes of acetone to 20 mL of lavage fluid. 2. Store overnight a 4' C to precipitate proteins. 3. Centrifuge at 1300g for 10 min. 4. Discard supernatant 5. Wash pellet with 10 mL of acetone, and centrifuge. 6. Discard acetone. 7. Dissolve the pellet in 1 mL of 0.2% trifluoroacetic acid,^ and store frozen

until ready for HPCE.

3. Ultrafiltration

The use of membrane filters in conjunction with centrifugation is a rapid way of purifying proteins, DNA, or RNA prior to separation. Salts, surfactants, biological debris, particles, and small molecules are easily removed. The technique does not require organic solvents and is faster than evaporation or lyophiliza-tion. Centrifugation increases the pressure to drive solvent and small molecules through the pores of the membrane. Membranes with molecular weight cutoffs of 3000, 10,000, 30,000, 50,000, and 100,000 are commercially available. The appropriate filter is selected based on the molecular weight of the target solute. Recovery of material is usually about 90% when the appropriate membrane is used. The salt concentration can be reduced by a factor of 100 if multiple cleanups are employed.

DNA-sequencing reaction products are best desalted by using spin columns^ (63). The columns should be hydrated for 30 min with 800 |iL of deionized water and washed with five aliquots of deionized water. The water each time is removed by centrifugation at 3000 rpm for 3 min. By running the sample through two columns, the salt concentration is reduced to 15 |LiM.

For the cleanup of PCR-amplified DNA, a 50-|iL volume is mixed with 2 mL of water. The mixture is purified using a Centricon (Millipore) with membranes that cut off MW 30,000 or 100,000, with centrifuging for 30 min at 5000g (33).

Following purification on a lysine-Sepharose column using an arginine gradient, recombinant human growth hormone is cleaned up using Centriprep (Millipore) followed by dialysis into 10 mM phosphate buffer, pH 2.5 (34).

4. Dialysis

Dialysis is a passive process that favors the transport of small molecules across a semipermeable membrane. Since small molecules have high diffusion coefficients, they encounter the membrane more frequently than do large molecules.

^TFA is a good protein solvent and does not interfere with electrophoresis at low pH. ^Centri-Spin (Princeton Separations, Adelphia, NJ).


Even if large molecules were to fit through the pores of the membrane, their rate of transport would be low because of their low diffusion coefficients. The process is facilitated when small volumes (e.g., 50-100 |LiL) are used. In this case, dialysis can be completed in less than 1 h. Dialysis is often used as part of the overall protein purification process (64).

In one example of DNA purification, proteins and excess salts were removed by phenol extraction and cold ethanol precipitation. The sample was then dia-lyzed for 2 days against citrate buffer to remove excess residual phenol and ethanol (43). Desalting of DNA can be performed by float dialysis (65). A drop of sample is placed on the CS 0.025-|Lim membrane, and dialysis is complete in 20 min. Over 80% of the chloride is removed.

10.6 MOBILITY AS A QUALITATIVE TOOL

Correction factors for chromatographic separations have been used for a long time. The best example is the use of retention indices for correcting intercolumn and instrumental variation in gas chromatography. For example, a solute with a retention index of 1250 elutes between n-Cu and n-Ci^ normal linear alkanes. In GC, the number of experimental variables is far fewer than in HPLC and HPCE. The primary variable in GC is the stationary phase. In HPLC, both the stationary phase and the mobile phase are important, and in HPCE, the carrier electrolyte dominates the experimental variables. Since the composition of the mobile phase or the carrier electrolyte is infinitely variable, the development of retention indices is difficult in HPLC or HPCE. There has been one report on the development of migration indices for HPCE (66), but it has not led anywhere.

Migration time is used for qualitative analysis, in an analogous fashion to retention time in chromatography. In HPCE, the migration time depends on both mobility and EOF Since EOF is more prone to drift than is mobility, it has been argued that mobility should be used as the qualitative parameter. Mobility should be independent of field strength and capillary length, but dependent on buffer composition and temperature. The solute's mobility is simply calculated by subtracting the electroosmotic mobility (determined with a neutral marker) from the apparent mobility.

The problem at hand is the reproducibility of the capillary inner wall. Bare silica capillaries exhibit capillary-to-capillary variation in EOF This results in "slow" and "fast" capillaries with regard to the measured migration times. Through the use of the combination of mobility data and spectra from diode array detection, HPCE has proven useful in forensic toxicological drug screening (9). The RSDs for mobility values for a 20-drug QC mixture were fractions of a percent over the course of weeks regardless of the capillary employed. The tremendous resolving power of CZE allows separations and drug identification not possible by HPLC. The system has been tested for proficiency and rugged-

10.7 Validation 4 4 5

ness, and according to John Hudson of the Royal Canadian Mounted Pohce, "CZE promises to compete with estabhshed chromatographic techniques as the screen of choice for forensic toxicologists."

10.7 VALIDATION

Much has changed over the last 20 years in the field of validation. Validation means confirming that a process performs as it is designed. Since the end product is a chemical entity such as a pharmaceutical substance, it is the entire process from synthesis to formulation to packing that must be validated. Analytical methods provide important information for this validation process. These methods must be validated in their own right. The degree of validation that is performed depends in part on whether regulatory agencies are involved and whether the separation is used to ensure health and/or safety.

The validation process consists of at least four steps (67) and possibly five:

1. Software validation 2. Hardware validation 3. Method validation 4. System suitability 5. Revalidation

The purpose of this section is to provide guidelines for items 2-5 using HPCE. These guidelines are nearly identical to those guidelines developed for HPLC. Since drug development is a worldwide process, the International Conference on Harmonization of Technical Requirements for the Registration of Pharmaceuticals has provided guidelines for the validation of methods.

As of March 1999, more than 50 HPCE methods have been validated and reported in the literature as this sampling indicates (18, 68-88).

A. HARDWARE VALIDATION

Hardware validation is performed to ensure that the equipment is working properly. While this is also monitored during method validation and routine operation, it is important to be able to distinguish a hardware problem that may require a service call from a problem in the method. Documentation should be established recording the serial numbers of the instruments, how the instrument was installed, and who performed the installation.

Most commercial systems provide kits for installation qualification (IQ) and operational qualification (OQ)-performance verification (PV). These can be employed to check detector noise, drift, linearity, wavelength accuracy, temperature stability, voltage stability, and other important parameters. If an analyst is


unable to reproduce a checkout sample following the manufacturer's protocol, either the hardware is in need of service or the analyst is in need of training. Most instruments also perform self-diagnostics to determine the integrity of electrical and mechanical components of the system.

Some instruments operate in a Good Laboratory Practice (GLP) mode. In one case, any postrun alteration of a data file results in a checksum failure. This is done to ensure the integrity of the data.

B. METHOD VALIDATION

This portion forms the core of this section, since most of the validation effort is applied here. To determine what needs to be validated, the type of analysis must first be determined. These procedures, from the ICH guidelines (67), are as follows:

1. Identification—simply identify a solute by its migration time and spectrum.

2. Quantitative—determine the impurities in a bulk drug. 3. Limit test—show that impurities are less than a specified amount. 4. Assay—quantitate the major substance.

Table 10.4 illustrates which parameters must be validated for a particular test. If US? assays are being validated, the guidelines change slightly depending on the nature of the tests. In Table 10.5, the meat of the validation process is described in outline form.

The key to controlling the vast amount of data that must be collected is ongoing documentation. Once the analyst falls behind, this task can be overwhelming.

Table 10.4 ICH Validation Characteristics versus Type of Analytical Procedure

Type of Analytical Procedure

Accuracy

Precision repeatability Interm. precision

Specificity

LOD

LOQ

Linearity

Range

Identification

No

No No

Yes

No

No

No

No

Impurity Testing

Quantitative

Yes

Yes Yes

Yes

No

Yes

Yes

Yes

Limit Tests

No

No No

Yes

Yes

No

No

No

Assay

Yes

Yes Yes

Yes

No

No

Yes

Yes

From reference (67).

10.7 Validation 4 4 7

Table 10.5 Method Validation

1. Define the purpose and scope of the method.

A. What analytes are to be separated?

B. What are the concentration ranges?

C. What are the limits of detection and quantitation?

D. Is the matrix defined?

E. Are there potential interferences?

E Are there regulatory requirements?

G. What is the required precision and accuracy?

H. Is a specific instrument required to perform the method?

1. Will the method be transferred among various laboratories?

J. What is the skill level requirements for the analyst?

2. Define the performance parameters and acceptance criteria.

3. Determine the critical performance characteristics of commercial instrumentation.

4. Check the quality of reference materials and reagents. Collect information on the chemical, physical, and toxicological properties of all solutes including degradation products.

5. Search the literature for prior art. Perform prevalidation experiments

6. Adjust performance parameters and/or acceptance criteria if necessary.

7. Validation experiments

A. Specificity using standards. The resolution should be greater than 2.5.

B. Linearity Run 5 standards covering the full working range. Inject each 3 times. Average the peak area, plot versus concentration, and calculate the linear regression. For impurity analysis, determine the rectilinearity the linear range of the trace impurity.

C. Precision. Inject a standard at 3 different concentrations, 5 times each. Calculate the relative standard deviation of the peak areas for each set of experiments. a. Run-to-run. b. Capillary-to-capillary c. Day-to-day (intermediate) for 15 days with 3 different analysts. d. Instrument-to-instrument.

D. Limit of detection (LOD). Inject a standard 3 times with a concentration close to the baseline noise. Average the signal height and baseline noise. The LOD is (3 x signal height)/baseline noise.

E. Limit of quantitation (LOQ). Prepare 6 standard solutions with amounts ranging from the expected LOQ to 20 times that amount. Inject all samples 6 times, and calculate the RSDs. The LOQ is usually defined when the RSD is 10% or when the signal-to-noise

to ratio is 10 (67).

E Accuracy. Spike a blank sample matrix with the solute at 3 different concentrations. Calculate the accuracy compared with the known values. This can be performed on a blind or double-blind basis to avoid bias in the results. All sample preparation steps

must be performed here.

G. Specificity. Use real samples and check peak purity using a diode array detector. It is also possible to fraction collect and analyze by another mode of HPCE.

(Continued)


Table 10.5 Method Validation (Continued)

H. Ruggedness. The degree of reproducibility of the results under a variety of experimental conditions expressed as %RSD. Among the variables to be considered are differences in (67): a. Laboratories and analysts. b. Capillaries and reagents. c. Instruments. d. Time periods.

1. Robustness. The capacity of a method to remain unaffected by small deliberate variations in method parameters (67). As a guideline, vary pH by ±0.5 pH units. All other parameters can be varied by ±10%. These include buffer concentration, capillary length, injection time, voltage, temperature, etc.

J. Cross-vahdation. If the new method is replacing an existing method such as HPLC, it is good practice to compare the two methods.

K. Solution stabihty. Determine the shelf life of all standards and reagents.

8. Develop standard operating procedures for routine performance of the method.

9. Define criteria for revalidation.

10. Define type and frequency of system suitability testing and/or quahty control checks of the acceptance criteria. Plan a course of action when the acceptance criteria are not met.

11. Fully document the validation experiments and results in the validation report.

A. Objective and scope.

B. Summary of method.

C. Solutes and matrix.

D. Specify reagents, electrolytes, reference standards, and quality control samples including purity, grade, and methods of preparation. Specify manufacturers and alternative suppliers.

E. Procedures for quality checks and standards and reagents.

E Safety precautions.

G. Implementation plan from R&D, methods transfer, and routine use.

H. Instrumental parameters.

I. Critical parameters found during robustness testing.

J. HPCE instrumentation and performance requirements.

K. Experimental conditions including sample preparation

L. Statistical procedures and sample calculations.

M. System suitability testing.

N. Representative electropherograms, spectra, and calibration curves.

O. Method acceptance limits and performance data.

P. Criteria for revalidation.

Q. Name of the method developer.

R. References.

S. Approval signatures, names, and titles.

Adapted with modifications from reference (90).

10.8 Troubleshooting 4 4 9

C. SYSTEM SUITABILITY

System suitability testing is used to ensure that the resolution and reproducibility of the system are up to the task to be performed. These tests verify the system as a whole. Among the parameters to be checked are plate count, resolution, migration time, and peak area reproducibility. A sample containing all of the expected components or the critical components is used as the test mix.

Unless otherwise specified, perform five runs when the %RSD is less than 2% and six runs when the %RSD is greater than 2%. This may have to be relaxed for long run times. The system suitability test should be performed at the beginning and at the end of a set of runs.

If a system suitability test fails, the analyst can make an adjustment in the method. If a method is changed, however, it must be revalidated. What constitutes a change and adjustment is a matter of semantics. For example, if a migration time window for a solute is specified to be from 9 to 10 min and the system suitability test gives a value of 10.1 min, it is reasonable to shorten the capillary by a few millimeters to bring the migration time into the specified range.

D. REVALIDATION

Upon methods transfer to a new laboratory, revalidation must be performed. A full validation is seldom required. A thorough system suitability test may prove sufficient when transferring to an experienced laboratory. Revalidation may also become necessary if sources of reagents, capillaries, or instruments are changed.

10.8 TROUBLESHOOTING

Table 10.6 (see following page ) contains an extensive troubleshooting guide. This guide can be used as an aid in methods development as well as for troubleshooting a previously working method. Most of these problems have been covered in the appropriate sections of this text.

450

Table 10.6 Troubleshooting

Chapter 10 Putting It All Together

Problem Cause and/or Solution

1. No peaks

A. Capillary problem

B. Voltage problem

C. Vial problem

D. Pneumatic problem

E. Detection problem

E Method error

G. Sample problem

H. Buffer problem

1. Service problem

2. Variable current


B. Injection related

C. Buffer related

D. Instrumentation

Replace or unplug capillary Broken capillary Misaligned capillary optical window Air bubble in capillary Solutes coat on capillary wall—use or replace coated capillary

Injected from incorrect side Incorrect polarity Voltage off

Empty sample vial Liquid level too low Vial cap missing or defective Air bubble in sample vial Solute coats vial wall

Check pressure or vacuum Turn on gas

Incorrect detector wavelength Detector lamp out Blocked optical window

Incorrect buffer or buffer position Short analysis time Incorrect vial position

Solutes degraded

Incorrect buffer Buffer has very high UV absorbance

Electrical, mechanical, or pneumatic failure—a service call

is required

Clogged capillary Broken capillary Air bubbles, degas buffer Cartridge incorrectly closed

Large injection (normal effect—reduce injection size if current drops to zero)

Incorrect buffer vial Empty or drained buffer vial Anolyte and catholyte are different

Coolant runs out Arcing, dirty electrodes Electrode not in vial Power supply problem—call service Current stable in one polarity, unstable the other way,

power supply problem—service call required

10.8 Troubleshooting 451

3. Poor peak area precision

A. Injection problem

B. Capillary problem

C. Sample problem

D. Buffer problem

E. Voltage problem

E Data system

4. Poor migration time precision


B. Buffer problem

C. Voltage problem

D. Temperature

Lengthen injection time Use internal standard Check injection pressure

Coating has deteriorated—replace capillary Wall effects—use coated capillary or additive Use acid or base intersample wash Increase buffer equilibration time Cartridge incorrectly closed

Viscosity variable between samples—dilute or perform sample preparation

Ionic strength variable between samples—see above Temperature variable between samples—allow longer

equilibration Sample depletion (electrokinetic injection)—replace sample Sample carryover—designate a wash station Evaporation—

Use closures Run fewer samples Lower sampler temperature

Buffer depletion—change buffers Contaminated buffers, change more frequently Evaporation problem Incorrect buffer Microbial growth

Use voltage ramp or insulate sample with buffer Contaminated electrodes

Inadequate sampling time Adjust integration parameters

Coating has deteriorated—replace capillary Wall effects—use coated capillary or additive Use acid or base intersample wash Increase buffer equilibration time Cartridge incorrectly closed

Buffer depletion—change buffers Contaminated buffers—change more frequently Evaporation problem Incorrect buffer

Contaminated electrodes

Ensure instrument operates at constant temperature

(Continued)


Table 10.6 Troubleshooting (Continued)

Problem Cause and/or Solution

5. Poor peak shape

A. Tailing

B. PeakspUtting

C. Broad peaks

6. Detection problems

A. Noisy or drifting baseline

B. Insufficient sensitivity

Ensure capillary is cut squarely Remove polyimide from capillary inlet Separate capillary and electrode Wall effects—use coatings or additives

Cracked capillary inlet Organic solvent in sample (MECC or CD)

Reduce injection size Lower ionic strength of injection buffer Siphoning—^balance fluid levels of buffer reservoirs Electrodispersion—

Lower sample concentration Increase BGE concentration Mobility match buffer to sample

Joule heating—run Ohms law plot

Replace detector lamp Filter and/or degas buffers Increase detector time constant Use low-UV absorbing buffer Tighten holder to reduce capillary vibration Realign capillary Clean electrodes Use a reference wavelength if possible

Optimize detector wavelength Increase sample concentration (stacking) Perform offline trace enrichment Increase capillary diameter Use extended path length capillary Use derivatization

REFERENCES

Weinberger, R., Albin, M. Quantitative Micellar Electrokinetic Capillary Chromatography: Linear Dynamic Range. J. Liq. Chromatogr., 1991; 14:953. Coors, C, Schulz, H.-G., Stache, E Development and Validation of a Bioanalytical Method for the Quantification of Diltiazem and Desacetyldiltiazen in Plasma by Capillary Zone Electrophoresis. J. Chromatogr, A, 1995; 717:235. Eap, C. B., Powell, K., Baumann, P Determination of the Enantiomers of Mianserin and Its Metabolites in Plasma by Capillary Electrophoresis after Liquid-Liquid Extraction and On-Column Sample Preconcentration. J. Chromatogr Sci, 1997; 35:315. Prunonosa, J., Obach, R., Diez-Cascon, A., Gouesclou, L. Determination of Cicletanine Enantiomers in Plasma by High-Performance Capillary Electrophoresis. J. Chromatogr, 1992; 574:127.

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5. Frost, M., Koehler, H., Blaschke, G. Determination of LSD in Blood by Capillary Electrophoresis with Laser-Induced Fluorescence Detection. J. Chromatogr., B: Biomed. Appl, 1997; 693:313.

6. Meier, P., Thormann, W. Determination of Thiopental in Human Serum and Plasma by High-Performance Capillary Electrophoresis-Micellar Electrokinetic Chromatography. J. Chromatogr., 1991; 559:505.

7. Makino, K., Goto, Y., Sueyasu, M., Futagami, K., Kataoka, Y., Oishi, R. Micellar Electrokinetic Capillary Chromatography for Therapeutic Drug Monitoring of Zonisamide. J. Chromatogr, B: Biomed. Appl, 1997; 695:417.

8. Lee, K.-J., Heo, G. S., Kim, N. J., Moon, D. C. Analysis of Anti-epileptic Drugs in Human Plasma Using Micellar Electrokinetic Capillary Chromatography. J. Chromatogr, 1992; 608:243.

9. Hudson, J. C , Golin, M., Malcome, M., Whiting, C. F Capillary Zone Electrophoresis in a Comprehensive Screen for Drugs of Forensic Interest in Whole Blood: An Update. J. Can. Soc. Forensic Scl, 1998; 31:1.

10. Nunez, M., Ferguson, J. E., Machacek, D., Jacob, G., Oda, R. R, Lawson, G. M., Landers, J. P Detection of Hypoglycemic Drugs in Human Urine Using Micellar Electrokinetic Chromatography Anal Chem, 1995; 67:3668.

11. Ashcroft, A. E., Major, H. J., Lowes, S., Wilson, 1. D. Identification of Non-steroidal Antiinflammatory Drugs and Their Metabolites in Solid Phase Extracts of Human Urine Using Capillary Electrophoresis-Mass Spectrometry. Anal Proc, 1995; 32:459.

12. Cavallaro, A., Piangerelli, V, Nerini, E, Cavalli, S., Reschiotto, C. Selective Determination of Aromatic Amines in Water Samples by Capillary Zone Electrophoresis and Sohd-Phase Extraction. J. Chromatogr, A, 1995; 709:361.

13. Steinmann, L., Thormann, W Toxicological Drug Screening and Confirmation by Electrokinetic Capillary Techniques: Concept of an Automated System. J. Capillary Electrophor, 1995; 2:81.

14. Wernly P, Thormann, W Drug of Abuse Confirmation in Human Urine Using Stepwise Solid-Phase Extraction and Micellar Electrokinetic Capillary Chromatography. Anal Chem., 1992; 64:2155.

15. Krynitsky, A. J. Determination of Sulfonylurea Herbicides in Water by Capillary Electrophoresis and by Liquid Chromatography/Mass Spectrometry. J. AOAC Int., 1997; 80:392.

16. Nguyen, A.-L., Luong, J. H. T. Separation and Determination of Polycyclic Aromatic Hydrocarbons by Solid Phase Microextraction/Cyclodextrin-Modified Capillary Electrophoresis. Anal Chem., 1997; 69:1726.

17. Riu, J., Schonsee, I., Barcelo, D. Determination of Sulfated Azo Dyes in Groundwater and Industrial Effluents by Automated Solid-Phase Extraction Followed by Capillary Elec-trophoresis/Mass Spectrometry. J. Mass Spectrom., 1998; 33:653.

18. Lucangioli, S. E., Rodriguez, V G., Otero, G. C. F, Vizioli, N. M., Carducci, C. N. Development and Validation of Capillary Electrophoresis Methods for Pharmaceutical Dissolution Assays. J. Capillary Electrophor, 1997; 4:27.

19. Wang, S.-R, Chang, C.-L. Determination of Parabens in Cosmetic Products by Supercritical Fluid Extraction and Capillary Zone Electrophoresis. Anal Chim. Acta, 1998; 377:85.

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21. Shihabi, Z. K., Hinsdale, M. E., Bleyer, A. J. Xanthine Analysis in Biological Fluids by Capillary Electrophoresis. J. Chromatogr, B: Biomed. Appl, 1995; 669:163.

22. Friedberg, M., Shihabi, Z. K. Ketoprofen Analysis in Serum by Capillary Electrophoresis. J. Chromatogr, B: Biomed. Appl, 1997; 695:193.

23. Ralston, P B., Strein, T. G. A Study of Deproteinization Methods for Subsequent Serum Analysis with Capillary Electrophoresis. Microchem.J., 1997; 55:270.


24. Shihabi, Z. K., Hinsdale, M. E. Analysis of Ibuprofen in Serum by Capillary Electrophoresis. J. Chromatogr., B: Biomed. Appl, 1996; 683:115.

25. Sczesny E, Hempel, G., Boos, J., Blaschke, G. Capillary Electrophoretic Drug Monitoring of Methotrexate and Leucovorin and Their Metabolites. J. Chromatogr., B: Biomed. Appl, 1998; 718:177.

26. Causse, E., Terrier, R., Champagne, S., Nertz, M., Valdiguie, P., Salvayre, R., Couderc, E Quantitation of Homocysteine in Human Plasma by Capillary Electrophoresis and Laser-Induced Fluorescence. J. Chromatogr, A, 1998; 817:181.

27. Hu, T, Zuo, H., Riley, CM., Stobaugh, J. E, Lunte, S. M. Determination of Of-Difluoromethy-lornithine in Blood by Microdialysis Sampling and Capillary Electrophoresis with UV Detection.;. Chromatogr, A, 1995; 716:381.

28. Hadwiger, M. E., Torchia, S. R., Park, S., Biggin, M. E., Lunte, C. E. Optimization of the Separation and Detection of the Enantiomers of Isoproterol in Microdialysis Samples by Cyclodex-trin-Modified Capillary Electrophoresis Using Electrochemical Detection. J. Chromatogr, B: Biomed. Appl, 1996; 681:241.

29. Hadwiger, M. E., Park, S., Torchia, S. R., Lunte, C. E. Simultaneous Determination of the Elimination Profiles of the Individual Enantiomers of Racemic Isoproterenol Using Capillary Electrophoresis and Microdialysis Sampling. J. Pharm. Biomed. Anal, 1997; 15:621.

30. Abubaker, M. A., Bissell, M. G., Petersen, J. R. Rapid Profiling of Clinically Relevant Steroids by Micellar Electrokinetic Capillary Chromatography. J. Capillary Electrophor, 1995; 2:105.

31. Kajiwara, H. Motif 2 in Adenosine Kinase Homologous Ginseng Polypeptide Showed Affinity to D-Ribose by Capillary Zone Electrophoresis and Surface Plasmon Resonance. J. Chromatogr, A, 1998; 817:173.

32. Cole, L. G., Kennedy, R. T. Selective Preconcentration for Capillary Zone Electrophoresis Using Protein G Immunoaffinity Capillary Chromatography. Electrophoresis, 1995; 16:549.

33. Schwartz, H. E., Ulfelder, K., Sunzeri, E J., Busch, M. R, Brownlee, R. G. Analysis of DNA Restriction Fragments and Polymerase Chain Reaction Products towards Detection of the AIDS (HIV-1) Virus in Blood. J. Chromatogr, 1991; 559:267.

34. Yim, K. W. Fractionation of the Human Recombinant Tissue Plasminogen Activator (rtPA) Glycoforms by High-Performance Capillary Zone Electrophoresis and Capillary Isoelectric Focusing. J. Chromatogr, 1991; 559:401.

35. Debernardi, S., Luzzana, M., De Bellis, G. Solid Phase Purification and SSCP Analysis of Amplified Genomic DNA by Capillary Electrophoresis. Front. Biosci. (Electronic Publication), 1996; 677:http://www.bioscience.org/bioscience/1996.

36. Figeys, D., Ducret, A., Yates, J. R., Ill, R., A. Protein Identification by Solid Phase Microex-traction-Capillary Zone Electrophoresis-Microelectrospray-Tandem Mass Spectrometry. Nat. Biotechnol, 1996; 14:1579.

37. Friedberg, M. A., Shihabi, Z. K. Urine Protein Analysis by Capillary Electrophoresis. Electrophoresis, 1997; 18:1836.

38. Figeys, D., Ducret, A., Aebersold, R. Identification of Proteins by Capillary Elec-trophoresis-Tandem Mass Spectrometry. Evaluation of an Online Solid-Phase Extraction Device. J. Chromatogr, A, 1997; 763:295.

39. Maier, M., Fritz, H., Gerster, M., Schewitz, J., Bayer, E. Quantitation of Phosphorothioate Oligonucleotides in Human Blood Plasma Using a Nanoparticle-Based Method for Sohd-Phase Extraction. Anal Chem., 1998; 70:2197.

40. Roche, M. E., Anderson, M. A., Oda, R. P, Riggs, B. L., Strausbauch, M. A., Okazaki, R., Wettstein, P. J., Landers, J. P Capillary Electrophoresis of Insulin-like Growth Factors: Enhanced Ultraviolet Detection Using Dynamically Coated Capillaries and Online Solid-Phase Extraction. Anal Biochem., 1998; 258:87.

41. Craig, D. B., Wong, J. C. Y., Polakowski, R., Dovichi, N. J. General Protease Assay Method Coupling Sohd-Phase Substrate Extraction and Capillary Electrophoresis. Anal Chem., 1998; 70:3824.

References 4 5 5

42. Belgrader, P., Devaney, J. M., Del Rio, S. A., Turner, K. A., Weaver, K. R., Marino, M. A. Automated Polymerase Chain Reaction Product Sample Preparation for Capillary Electrophoresis Analysis. J. Chromatogr., B: Biomed. Appl, 1996; 683:109.

43. Jacquier, J.-C, Gorelov, A. V, McLoughlin, D. M., Dawson, K. A. Capillary Electrophoretic Study of the Complex Formation Between DNA and Cationic Surfactants. J. Chromatogr., A, 1998; 817:263.

44. Heo, Y. J., Kim, S. Y., Kim, E., Lee, K.-J. Quaternary Structural Analysis of Nucleoside Diphosphate Kinases Using Capillary Electrophoresis. J. Chromatogr., A, 1997; 781:251.

45. Piggee, C. A., Muth, J., Carrhilo, E., Karger, B. L. Capillary Electrophoresis for the Detection of Known Point Mutations by Single-Nucleotide Primer Extension and Laser-Induced Fluorescence. J. Chromatogr, A, 1997; 781:367.

46. Bourque, A. J., Cohen, A. S. Quantitative Analysis of Phosphorothioate Oligonucleotides in Biological Fluids Using Direct Injection Fast Anion-Exchange Chromatography and Capillary Gel Electrophoresis. J. Chromatogr, B: Biomed. Apph, 1994; 662:343.

47. Gelfi, C, Orso, A., Righetti, P. G. Capillary Zone Electrophoresis in Polymer Networks of Polymerase Chain Reaction-Amplified Oligonucleotides: The Case of Congenital Adrenal Hyperplasia. J. Chromatogr, B: Biomed. Appl, 1994; 657:201.

48. Klyushnichenko, V., Kula, M.-R. Rapid SDS Gel Electrophoretic Analysis of Proteins. J. Capillary Electrophor, 1997; 4:61.

49. Wang, Y., Wallin, J. M., Ju, J., Sensabaugh, G. E, Mathies, R. A. High-Resolution Capillary Array Electrophoretic Sizing of Multiplexed Short Tandem Repeat Loci Using Energy-Transfer Fluorescent Primers. Electrophoresis, 1996; 17:1485.

50. Nathakarnkitool, S., Oefner, P J., Bartsch, G., Chin, M. A., Bonn, G. K. High-Resolution Capillary Electrophoretic Analysis of DNA in Free Solution. Electrophoresis, 1992; 13:18.

51. Causse, E., Simeon, N., Nertz, M., Salvayre, R., Bayard, F, Valdiguie, P., Couderc, E Quantitation of RT-PCR Products of bFGF-mRNA by Capillary Electrophoresis and Laser-Induced Fluorescence. J. Capillary Electrophor, 1997; 4:77.

52. Marsh, M., Tu, O., Dolnik, V, Roach, D., Solomon, N., Bechtol, K., Smietana, P, Wang, L., Li, X., Cartwright, P, Marks, A., Barker, D., Harris, D., Bashkin, J. High-Throughput DNA Sequencing on a Capillary Array Electrophoresis System. J. Capillary Electrophor, 1997; 4:83.

53. Cheng, J., Kasuga, T., Watson, N. D., Mitchelson, K. R. Enhanced Single-Stranded DNA Conformation Polymorphism Analysis by Entangled Solution Capillary Electrophoresis. J. Capillary Electrophor, 1995; 2:24.

54. Gurley, L. R., Buchanon, J. S., London, J. E., Stavert, D. M., Lehnert, B. E. High-Performance Capillary Electrophoresis of Proteins from the Fluid Lining of the Lungs of Rats Exposed to Perfluoroisobutylene. J. Chromatogr, 1991; 559:411.

55. Castaneda Penalvo, G., Kelly, M., Maillois, H., Fabre, H. Evaluation of Capillary Zone Electrophoresis and Micellar Electrokinetic Capillary Chromatography with Direct Injection of Plasma for the Determination of Cefotaxime and Its Metabohte. Anal. Chem., 1997; 69:1364.

56. von Herrer, F, Tanner, R., Theurillat, R., Thormann, W Determination of Fluconazole in Human Plasma by Micellar Electrokinetic Capillary Chromatography with Detection at 190 nm. J. Chromatogr, A, 1996; 745:165.

57. Nishi, H., Fukuyama, T, Matsuo, M. Separation and Determination of Aspoxicillin in Human Plasma by Micellar Electrokinetic Chromatography with Direct Sample Injection. J. Chromatogr, 1990; 515:245.

58. Thormann, W, Minger, A., Molteni, S., Caslavska, J., Gebauer, P Determination of Substituted Purines in Body Fluids by Micellar Electrokinetic Capillary Chromatography with Direct Sample Injection. J. Chromatogr, 1992; 593:275.

59. Shihabi, Z. K. Peptide Stacking by Acetonitrile-Salt Mixtures for Capillary Zone Electrophoresis. J. Chromatogr, A, 1996; 744:231.

60. Shihabi, Z. K. Sample Stacking by Acetonitrile-Salt Mixtures. J. Capillary Electrophor, 1995; 2:267.


61. Shihabi, Z. K. Stacking of Weakly Cationic Compounds by Acetonitrile for Capillary Electrophoresis. J. Chromatogr., A, 1998; 817:25.


63. Ruiz-Martinez, M. C , Salas-Solano, O., Carrilko, E., Kolter, L., Karger, B. L. A Sample Purification Method for Rugged and High-Performance DNA Sequencing by Capillary Electrophoresis Using Replaceable Polymer Solutions. A. Development of the Cleanup Protocol. Anal. Chan., 1998; 70:1516.

64. Reif, O. W, Freitag, R. Control of the Cultivation Process of Antithrombin III and Its Characterization by Capillary Electrophoresis. J. Chromatogr, A, 1994; 680:383.

65. Williams, R E., Marino, M. A., Del Rio, S. A., Turni, L. A., Devaney J. M. Analysis of DNA Restriction Fragments and Polymerase Chain Reaction Products by Capillary Electrophoresis. J. Chromatogr, A, 1994; 680:525.

66. Lee, T. T., Yeung, E. S. Facihtating Data Transfer and Improving Precision in Capillary Electrophoresis with Migration Indices. Anal. Chem., 1991; 63:2842.

67. Swartz, M. E., Krull, I. S., Analytical Method Development and Validation. 1998, Marcel Dekker. 92.

68. Altria, K. D., Chanter, Y. L. Vahdation of a Capillary Electrophoresis Method for the Determination of a Quinolone Antibiotic and Its Related Impurities. J. Chromatogr, 1993; 652:459.

69. Altria, K. D., Clayton, N. G., Harden, R. C, Makwana, J. Inter-company Cross Validation Exercise on Capillary Electrophoresis. Quantitative Determination of Drug Counter-ion Levels. Chromatographia, 1995; 40:47.

70. Altria, K. D., Rudd, D. R. An Overview of Method Validation and System Suitability Aspects in Capillary Electrophoresis. Chromatographia, 1995; 41:325.

71. Arellano, M., Andrianary J., Dedieu, F, Couderc, F, Puig, P. Method Development and Validation for the Simultaneous Determination of Organic and Inorganic Acids by Capillary Zone Electrophoresis. J. Chromatogr, A, 1997; 765:321.

72. Berzas, N. J. J., Guiberteau Cabanillas, C , Contento Salcedo, A. M. Method Development and Validation for the Simultaneous Determination of Dyes in Foodstuffs by Capillary Electrophoresis. Anal Chim. Acta, 1998; 378:65.

73. Cherkaoui, S., Mateus, L., Christen, P, Veuthey, J.-L. Development and Validation of a Capillary Zone Electrophoresis Method for the Determination of Atropine, Homoatropine and Scopalamine in Ophthalmic Solutions. J. Chromatogr, B: Biomed. Appl, 1997; 696:283.

74. Dimo-Simonin, N., Grange, F, Kratzer, A., Brandt-Casadevall, C, Mangrin, P. Forensic Validation of the Short Tandem Repeat HUMACTBP2 Using Capillary Electrophoresis. Electrophoresis, 1998; 19:256.

75. Emaldi, P, Fapanni, S., Baldini, A. Vahdation of a Capillary Electrophoresis Method for the Determination of Cephradine and Its Related Impurities. J. Chromatogr, A, 1995; 711:339.

76. Fabre, H., Blanchin, M. D., Julien, E., Segonds, C , Mandrou, B., Bosc, N. Validation of a Capillary Electrophoresis Procedure for the Determination of Calcium in Calcium Acamprosate. J. Chromatogr, A, 1997; 772:265.

77. Hunt, G., Hotaling, T, Chen, A. B. Vahdation of a Capillary Isoelectric Focusing Method for the Recombinant Monoclonal Antibody C2B8. J. Chromatogr, A, 1998; 800:355.

78. Kelly, M. A., Altria, K. D., Grace, C , Clark, B. J. Optimization, Vahdation and Application of a Capillary Electrophoresis Method for the Determination of Ranitidine and Related Substances. J. Chromatogr, A, 1998; 798:297.

79. Kohn, A. Automated Method Validation and System Suitabihty Testing in HPLC and CE. Am. Biotech. Lah, 1994; 12:44.

80. LaFountain, M., M., S., Cormier, J., Buel, E. Validation of Capillary Electrophoresis for Analysis of the X-Y Homologous Amelogenin Gene. J. Forensic Set, 1998; 43:1188.

81. Moorhouse, K. G., Rickel, C. A., Chen, A. B. Electrophoretic Separation of Recombinant Tissue-Type Plasminogen Activator Glycoforms: Validation Issues for Capillary Isoelectric Focusing Methods. Electrophoresis, 1996; 17:423.

References 457

82. Castaneda Penalvo, G., Julien, E., Fabre, H. Cross Validation of Capillary Electrophoresis and High-Performance Liquid Chromatography for Cefotaxime and Related Impurities. Chro-matographia, 1996; 42:159.

83. Ross, G. A. Instrumental Validation in Capillary Electrophoresis and Checkpoints for Method Validation. Accredit. Qual. Assur, 1997; 2:275.

84. Saenger-Van De Griend, C. E., Wahlstroem, H., Groeningsson, K., Widahl-Naesman, M. A Chiral Capillary Electrophoresis Method for Ropivacain Hydrochloride in Pharmaceutical Formulations: Vahdation and Comparison with Chiral Liquid Chromatography J. Pharm. Bio-med.Anal., 1997; 15:1051.

85. Shah, P A., Quinones, L. Validation of a Micellar Electrokinetic Capillary Chromatography (MECC) Method for the Determination of p-Toluenesulfonic Acid Impurity in a Pharmaceutical Intermediate. J. Liq. Chromatogr., 1995; 18:1349.

86. Swartz, M. E., Mazzeo, J. R., Grover, E. R., Brown, P R. Validation of Enantiomeric Separations by Micellar Electrokinetic Capillary Chromatography Using Synthetic Chiral Surfactants. J. Chromatogr., A, 1996; 735:303.

87. Wielgos, T., Turner, P, Havel, K. Validation of Analytical Capillary Electrophoresis Methods for Use in a Regulated Environment. J. Capillary Electrophor, 1997; 4:273.

88. Yesilada, A., Tozkoparan, B., Gokhan, N., Oner, L., Ertan, M. Development and Validation of a Capillary Electrophoretic Method for the Determination of a Degradation Product in Nap-hazoline HCl Bulk Drug Substance. J. Liq. Chromatogr Related Technol, 1998; 21:2575.

89. Altria, K. D., Harden, R. C, Hart, M., Hevizi, J. Inter-company Cross-Validation Exercise on Capillary Electrophoresis. I. Chiral Analysis of Clenbuterol. J. Chromatogr, 1993; 641:147.

90. Huber, L. Validation of HPLC Methods. BioPharm, 1999; 12:64.

INDEX

Absorption detection bandbroadening, 370-372 diode array, 372-373 extended pathlength, 375-379 indirect detection, see Indirect

detection wavelength optimization,

375, 376 Acids, aromatic, 120 Affinity capillary electrophoresis,

194, 195 (table), 196 Amino acids,

chiral, 180, 181, 191-192 indirect detection, 103 MECC, 167, 388, 389 nonaqueous, 120 stacking of, 342, 350

Aminoglycosides, 96 Aminomethylphenols, 235 Amino sugars, 121 Amphetamines,

chiral, 180 CZE, 96 MECC, 167 nonaqueous, 120

Amphotericin, 167 Angiotensins,

CEC, 310 MECC, 167

Anions, small, 102-103, 110-111, 120

Anthracyclines, 96 Antihistamines, 120 Antisense DNA, see DNA Antistacking, 27, 340-343 Antithrombin 111, 93, 235 Arginase isoforms, 93 Ascorbic acid, 96 Aspartame, 96 Aspoxicillin, 438

Atrizine, 167 Atrolactic acid, 180 Atropine, 167

Background electrolyte, 1 Bandbroadening, 64, 66-69,

66 (table) antistacking, 66, 340-343 detection, 370-372 diffusional, 41 dispersive transport, 7 electrodispersion, 67-68,

431-432 hydrodynamic flow, 40 injection, 321-323, 331-332 joule heating, 43-47 mass transfer, 39 siphoning, 332 wall effects, 78-83

Barbiturates, 8-9, 167, 440 Benzene and derivatives, 310 Benzodiazepines, 167 Biphenyls, 310 Bradykinin, 96 Buffers, 5^58, 55 (table), 56 (table)

conductivity, 26 depletion, 26-27, 230, 424 preparation, 57-58 selection of, 56-57

Buffer additives, 59-61 (table) cationic surfactants, 86-87 diaminopropane, 85 non-ionic surfactants, 89-91

Buserelin acetate, 96

Cannabinoids, 310 Capillaries

coatings, 64-65 (table) diameter, 45-46 dynamic coatings, 86-89

fused-silica, 60-61 GC capillaries, 89 length, 30, 52-54 preparation, 62-63 Storage, 63-64 volume, 321 (table) wash procedures, 63, 425

Capillary electrochromatography, 10

applications, 309, 310-311 (table), 312-313

bubble formation, 297 chiral, 296, 314 column equilibration, 305 column packing, 303-304, 306 eddy diffusion, 303-303 efficiency 301-303, 305 (table) electroosmotic flow, 299 (table) flow rate, 294 (table) frits, 297 injection, 306 instrumentation, 297 method development,

306-308 microfluidic devices, 313, 316 nominal flow rates, 294 open-tubular, 295-296 packed, 297-298 packing materials, 299 replaceable media, 298 tailing, 297

Capillary electrophoresis history 11-12, 13 (table), 14 instrumentation, 15 (table),

16-18, hterature, 19-20 modes of, 20-21 mode selection, 423 (table) optimization, 426 (table) properties, 9-10

459

460 Index

Capillary electrophoresis (continued)

textbooks, 20 (table) Capillary isoelectric focusing

additives for hydrophobic proteins, 221-222


buffer depletion, 230 capillaries, 215-217

conditioning, 222 carrier ampholytes, 210-211

narrow range, 220-221 UV background, 218-219

detection, 232-234 focusing, 210-211, 225-226 injection, 224-225 internal standards, 217 (table),

218 mechanism, 210-211 mobilization, 210, 226-230 pH gradient formation,

212-213 preparation of methylcellulose

solution, 218 protocols, 223 (table) resolving power, 214-215 saU effects, 230-231, 233 sample preparation validation, 240

Capillary zone electrophoresis applications

capillary ion analysis, 99-102, 103 (table), 104-113

carbohydrates, 119-120, 121 (table), 122-123

proteins, 93-94 (table) small molecules, peptides,

96-97 (table) bandbroadening, 64, 66-69 efficiency, 39-43

effect of capillary length, 52-54

injection, see Injection mobility plot, 74-78 nonaqueous electrophoresis,

116-119, 120 (table) solvents for, 119 (table)

resolution, 41-42 effect of capillary length,

52-54 secondary equilibrium, 95,

98-99 separation strategies, 90-92,

95, 98-99 wall effects, see Wall effects

Carbohydrates, aliphatic, 103 aromatic, 96 carboxylic acids, indirect detection, 103 organic, 96

Cardiac glycosides, 167 Caridopa, 180 Casein phosphopeptide, 96 Catalyse, 273 Cations, metal, 103, 108-109, 329

equivalent ionic conductance, 108

physical properties, 29 (table) Cefixime, 96 Cephalosporins, 167 Chiral recognition, 87-88,179,

180-181 (table), 182-194, 296 reagents for,

bile salts, 192-193 crown ethers, 188-189 cyclodextrins, 182-188 macrocyclic antibiotics,

189-191,314 metal-ion complex, 179, 182 oligosaccharides, 191-192 surfactants, 182

Chloramphenicol, 185 Chlorophylls, 167 Chlothalidone, 296,310 Choline esters, aromatic, 167 Cocaine, 180 Collagen, 93 Conductivity detection, 112-113 Corticosteroids, 161, 165-166 Coumarin, 180 Chorionic gonadotropin

glycoforms, 93 Creatinine, 103 Critical micelle concentration,

141-142 Cyclodextrins, 161-163 (table),

164-166 Cyclodextrin-MECC, 164-166 Cytochrome c, 401

Deoxyribonucleosides, 167 Deoxyribonucleotides, 97, 167 Derivatization,

chiral, 193-194 on-capillary 390 post-capillary, 390-392 pre-capillary, 385-390 reagents for, 386 (table), 387

(table) Detection

absorption detection, see Absorption detection

concentration limit of detection, 368, 374-375

fluorescence, see Fluorescence detection

indirect detection, see Indirect detection

mass limit of detection, 368 mass spectrometry, see Mass

spectrometry peak area normalization,

366-367 time constant, 371 (table), 372 types of detectors, 369 (table)

Dialysis, see Sample preparation Diflunisal, 372 Diltiazem, 180, 193 Dispersive transport, 7 Diffusion, 41-42, 81 (table) DNA

adducts, 310 antisense, 266, 268-271

bases, 168 genetic analysis, 266, 281-283 hybridization products, 266, 276 intercalators, 261, 281, 390 microsatellites, 266 oligonucleotides, 252, 266,

274-275 comparison with HPLC,

275 (table) plasmids, 276, 283 restriction digest, 256, 263,

267, 278-280, 285 sample preparation, 279,

442-444 short tandem repeats, 266,

280-282 sequencing, 3, 252, 266,

276-278 sizing ladder, 257

DNA Sequencing, see DNA

Ecdysteroids, 168 Electromigration dispersion,

see Bandbroadening Electroendoosmosis

CEC, 299-300 capillary surface, 31-33 control of, 38 direction of, 34-35, 43, 102 effect of

buffer concentration, 35-37 capillary surface, 31-33 field strength, 37 organic solvents, 37-38 pH, 34-36 temperature, 58-59, 424 viscosity, 37

hysteresis, 34, 36 measurement of, 32-34 reversal of, 38, 86-87 smoluchowski equation, 32 suppression of, 38

Electroneutrality, 26 Enkephalins, 96 Enzyme assays, 390 Epinephrine, 88, 181 Ergot alkaloids, 181 Estrogens, 168 Etoposide phosphate, 96 Explosives, 168

Fatty acids, short chain, 103 Fatty acids, 168 Ferguson plot, 278-280 Field-amplified injection,

see Stacking Field strength, 31 Flavonoids, 168 Flavonol glycosides, 168 Fluorescence detection

basic concepts, 379-380 DNA Sequencing, 384 optimization, 380-383 laser-induced, 383-385

laser wavelengths, 384 (table) Fluticasone propionate, 311 Fraction collection, 405-408

Index 461

Gangliosides, 121 Gas chromatography, 4 Genetic analysis, see DNA Granulocyte macrophage colony

stimulating factor, 235 Henderson-Hasselbalch equation,

76 (table), 77 Hemoglobins, 235, 237-238 Herbicides, 168, 180 Heroin, 175-177 Heroin impurities, 45-46 Histones, 93 Human bone morphogenetic

protein glycoforms, 93 Human growth hormone,

isoforms, 93 post translational

modifications, 93 precursor, 93 tryptic digest, HPLC, 115 tryptic digest, CZE, 116-118

Human immunodeficiency provirus DNA, 279

Human rhino virus, 236 Hydrocarbons, aromatic, 120 Hydrochlorothiazide, 168

Imipramine, 120 Immunoassay, 389-390 Immunoglobulins, 93 Indirect detection, 103 (table),

104-107 mobility matching, 104-106 reagent selection, 106 (table)

Indirect fluorescence detection. 111

Injection, bandbroadening from, 321-323

(table), 331-332 capillary volume, 321 (table) flowrate, 326 (table), 327 electrokinetic, 327, 329-330

mobility bias, 329 ionic strength bias, 278

(table), 329-330 linearity, 330

hydrodynamic, 325-327 performing a run, 323-324 poiseuille equation, 325 stacking, see Stacking troubleshooting, 331 ubiquitous (spontaneous)

injection, 332 viscosity effects, 326, 425 volume, 326

Ink, fountain pen, 96 Insulin, 93 Insulin receptor peptide, 168 Interleukin II, 93 Internal standard, 425 lohexol, 440 Isoflavones, 96 Isoproterenol, 181 Isotachophoresis, see Transient

isotachophoresis Isradipin, 313

Joule heating, 43-47

a-Lactoglobulin, 60 Laser-induced fluorescence, see

Fluorescence detection Laminar flow, 39 Lanthanides, 103 Leucovorin, 181 Limit of detection, 4, 7, 368

indirect detection, 107 Lipoproteins, 93 Liquid chromatography, 5-8,

10,16 oligonucleotides, 275 resistance to mass transfer, 39

Lysine, 76

Macrolide antibiotics, 168 Mandelic acid, 181 Mass spectrometry

electrospray 395-403 buffers, 397 interfaces, 400-403 ion formation, 396 molecular weight calculation,

398-399 (table) sensitivity 400 (table)

fast-atom bombardment, 403-405

nonaqueous, 120 HPCE techniques, 395 (table)

Mefentidine, 120 Mefloquine, 181 Membrane proteins, 93 Metal chelates

acetylacetone, 168 PAR, 168

Metallothionein isoforms, 93, 168 Methotrexate, 96 Micellar electrokinetic capillary

chromatography, 6-7, 12 applications, 166, 167-169

(table), 170-176 capacity factor, 144-145 electroosmotic flow, 152 elution order, 146-149 mechanism, 143-145 measurement of to

and tmc, 154 methods development,

176-178 optimization

organic solvents, 154-156 pH, 151-153 surfactant concentration,

148-150 urea, 157

resolution, 145 reversed-polarity 152-153 stacking, 336-337, 345-346 surfactants, 158-160 (table)

Micelles, 141-143 Micro-liquid chromatography,

5, 10, Micromachined electrophoretic

devices, 11-12 Microsatellites, see DNA Migration time, 28 Milk proteins, 94 Mobihty 1, 28-31, 74

apparent, 32-33 correction, 444-445 effect of,

buffer concentration, 50-53 pH, 75-79 temperature, 58-60

direction of, 43 matching, 105-106

Monoclonal antibody, CIEF, 236 doxirubicin conjugate, 168

Monosaccharides, 121 Morphine-3- glucuronide, 168 Motilin peptides, 96 Mucin glycoforms, 94 Myoglobins, 397

Naphthalene sulfonates, 311, 312 Nitrate, 97 Nitro toluenes, 168 Nonaqueous electrophoresis,

116-119 Non-steroidal antiinflammatory

drugs, chiral, 180, 181 CZE, 97 MECC, 147-149, 169

Nuclease A variants, 94 Nucleosides, 97, 169 Nucleoside diphosphate

kinases, 94 Nucleosides, 101, 378

Ohm's law, 26 Ohm's law plot, 47-30 Oligonucleotides,

HPLC, 275 MECC, 169 size separation, 251, 266, 274

Oligosaccharides, ANTS labeled, 121 APTS labeled, 121 dextran, 121, 124 heparin, 121 sialio, 121, 169

Ovalbumin glycoforms, 94

Parabens, MECC, 153 CEC, 311

Parabolic flow, see Laminar flow Peak tailing,

cracked capillary inlet, 333 electrodispersion,

see Bandbroadening wall effects, see Wall effects

Peak sphtting, 334 Penicillins, 169 Peptides, misc.,

CEC, 311 chiral, 181, 187-188 CZE, 51-52, 59, 79 low-UV detection of, 376 mapping, 112-113, 114 (table),

115-118,382,392 MECC, 146 stacking of, 345

462 Index

Pesticides carbamate, 169 urea, 169

Plasmid DNA, see DNA Polycyclic aromatic hydrocarbons,

MECC, 169 CEC, 304, 311,314

Polymyxins, 169 Polyphosphate, 103 Polysaccharides, 121 Porphyrins

metal complex, 169 urinary, 169, 170-175,

336-337, 358-359 Phospholipids, 103 Phosphonic acids, alkyl, 103 Phytate, 103 Polymerase chain reaction

products, 3-5, 279 Polymer network, see Size

separations Post-capillary derivatization,

see Derivatization Pre-capillary derivatization,

see Derivatization Procanamide, 97 Propranolol, 181 Proteins, misc.,

CZE, 85, 90-91 serum, 123, 125 sample preparation, 442-444 size separation, 252, 259, 260,

267, 270-274 SDS-protein complex

preparation, 271-272 Purines, 169

Quantitative analysis area percent, 431 data sampling rate, 427 external standards, 428 internal standards, 430 linearity 330, 431-433, 436 peak areas, 427

Quinagohde, 187

Rattlesnake mycotoxins, 94 Restriction digest, see DNA Ribonucleases, 239 Ribonucleotides, 97 RNA, 267, 283-284

Salbutamol, 181 Sample depletion, 334 Sample preparation

basic principles, 434, 436 biological fluids, 437-441

direct injection, 437-438 protein precipitation, 439

liquid-liquid extraction, 439 solid-phase extraction,

439, 441 dialysis, 443-442 precipitation, 442-443 ultrafiltration, 279, 443

Sodium dodecyl sulfate, determination, 103

Separative transport, 6 Serum proteins, 94 Short tandem repeats, see DNA Size separations


agarose, 250-251 capillaries, 255 crosslinked polyacrylamide,

249-250, 252 (table) detection

LIE, Uy 261

intercalators, 261, 263-264 materials for, 254, 257-262 methods development, 268-271 polymer networks, 252-262

commercial kits, 262 (table) concentration, 253 field strength, 254-255 injection, 254 materials, 254 (table) temperature, 254-255

pulsed-field, 247, 284-286 separation mechanism,

247-248 Slab-gel electrophoresis, 1-4, 6,10,

isoelectric focusing, 209 size separations, 245-246

Spontaneous injection,

see Injection Spontaneous peaks, 332 Stacking, 26, 335

acetonitrile-salt mediated, 344 field-amplified injection,

351-352, 353 (table), 354 indirect detection, 109-110 ionic-strength-mediated,

336-340 isotachophoresis, see Transient

isotachophoresis LC-based, 359-360 MECC, 345-346 membrane-based, 359-360 neutral solutes, 345-347 pH mediated, 343-345 whole-capillary injection,

349-350 Steroids, 311,347 Stokes law, 28 Structural isomers, 100

Substance P fragments, 328 Sulfonamides, 97 Sugar, phosphorylated, 121

Tailing, see Peak tailing Tamoxifin, 120 Terbutaline, 183 Thalidomide, 314 Theophylline, 169, 440 Theoretical plates, 7, 41, 42 (table) Tioconazole, 181 Tissue plasminogen activator,

CIEF, 216, 219, 236 validation, 240-241

Transferrins, 94 Transient isotachophoresis,

354-359 buffers for, 357 (table),

358 (table) mechanism, 354-355 practical advise, 358-359

Triazine herbicides, 236 Tricyclic antidepressants, 169 Tropane alkaloids, 120 Troubleshooting, 450-452 (table) Tumor necrosis factor, 94

Ubiquitous injection, see Injection

Ultrafiltration, see Sample preparation

UV detection, see Detection

Validation CIEF, 240 hardware, 445-446 intercompany, 429 (table),

430 (table) method, 446, 447-448 (table),

449 revalidation, 449 system suitability, 449

Velocity, electrophoretic, 31 Verapamil, 181 Vitamins

fat soluble, 120 MECC of, 150-151, 167, 169

Wall effects, 78-91,424 ion-pairing, 80 protein recovery, 83 (table) random walk model, 81-82 reduction of, 83-89

Warfarin, 181 Whey proteins, 94

Xanthines, 169

Zeta potential, 31, 35

Practical capillary electrophoresis

Education

Transcript of Practical capillary electrophoresis