Acid and base hydrolysis of lipid A fromEnterobacter agglomerans as monitored by electrospray...

JOURNAL OF MASS SPECTROMETRY, VOL. 31, 138-149 (1996)

Acid and Base Hydrolysis of Lipid A from Enterobacter agglomerans as Monitored by Electrospray Ionization Mass Spectrometry : Pertinence to Detoxification Mechanisms

Yan Wang and Richard B. Cole*

Department of Chemistry, University of New Orleans, Lakefront, New Orleans, Louisiana 70148, USA

Lipopolysaccharides (LPS), which are endotoxins found in the cell wall of Gram-negative bacteria, are common components of organic dusts that cause or contribute to symptoms associated with organic dust diseases. The lipid A subgroup within LPS is believed to be responsible for the toxicity. Acid and base treatments, which can be effective detoxification methods, were performed on lipid A from Enterobucter ugglomeruns (EA), a bacterium commonly found in field cotton. Negative-ion electrospray ionization mass spectrometry was employed to charac- terize the post-treatment structural changes to lipid A. Acid treatment (1% acetic acid, 1OO'C) hydrolyzed the ester side-chains of lipid A. I t was found that the ester-linked palmitoyl group was the most labile to acid hydrolysis. Hydrolysis of the palmitoyl moiety conformed to pseudo-first-order chemical reaction kinetics with a rate constant for decomposition of heptaacyl-lipid A from Enterobucfer uggfomeruns of -3.3 x min-'. An order of lability of lipid A acyl side-chains to acid hydrolysis was also deduced: R,' (palmitoyl) > R,' (myristoyl or hydroxymyristoyl) > R, (hydroxymyristoyl at position 3) > R, (oxymyristoyl group at position 3 ) > R,' (lauroyl). Base treatment (0.05 M NaOH in 95% EtOH, 65'C) was shown to be more effective at cleaving ester-linked side-chains. In addition, mass spectral evidence suggests that opening of the pyranose rings of the disaccharide backbone of lipid A and/or removal of the phosphoryl groups may be occurring during base treatment. This study sheds light on mechanistic aspects of treatment procedures leading to the detoxification of endotoxins.

KEYWORDS: electrospray; mass spectrometry; lipid A; Enterobacter agglomerans; detoxification

INTRODUCTION

Lipopolysaccharides (LPS) are endotoxins produced by all Gram-negative bacteria. They elicit a variety of biological, medical and immunological activities such as induction of fever, change in the white blood cell count, diarrhea or even shock and lethal toxicity."' Because of these important properties, the study of the relationship between LPS structure and its toxicity has been an area of intensive i n v e ~ t i g a t i o n . ~ ~ ~ Despite their considerable structural diversity, all LPS are built upon three cova- lently interlinked distinct structural regions: the 0- specific chain, the core oligosaccharide and the lipid A component,6 as shown in Scheme 1. The O-specific chain and the oligosaccharide parts are hydrophilic, whereas the lipid A moiety is hydrophobic. The poly-

[O-Specific Chain] - [Outer Core] - [Inner Core - (KDO),] - [Lipid A]

Core Region 1 Scheme 1. Schematic representation of the components con- tained in enterobacterial lipopolysaccharides.

* Author to whom correspondence should be addressed.

O H

R'i = C=O or C=O ; c=o I I I

I y 2 ( y 4 2 ) 1 2 CH2 I p H O - y H CH3

a HO-CH

10 ( y 2 ) l o

CH3 CH3

Figure 1. Structure of monophosphorylheptaacyl-lipid A from Enterobacter agglomerans .

CCC 1076-5 174/96/020138-12 0 1996 by John Wiley & Sons, Ltd.

Received 22 June 1995 Accepted 3 October 1995

HYDROLYSIS OF LIPID A AS MONITORED BY ESMS 139

saccharide and the lipid A portions are joined by a covalent linkage between KDO (2-keto-3-deoxyoctinic acid) and the 6'-position of the glucosamine backbone of lipid A (Fig. 1). The acid lability of this bond permits the use of acid hydrolysis to divide the LPS into hydrophilic and hydrophobic components; the latter precipi- tates out of aqueous solution.

Previous toxicity studies have established that the toxic activity of such endotoxins originates in the lipid A p ~ r t i o n . ~ - ~ Accordingly, over the last two decades, studies have focused on identifying the structure of lipid As from different bacterial species, while probing the relationship with toxicity.'*-12 Only fairly recently, and owing to the continuing development and improvement of analytical techniques, has the entire structure of LPS (especially the core and lipid A regions) from certain bacteria been fully described.' 3314

Mass spectrometry, a well established tool for the structural characterization of organic compounds, has played an indispensable role in the ability to determine lipid A structures. Many different mass spectrometric ionization techniques have been applied to the analysis of lipid A from various types of bacteria. Fast atom bombardment (FAB) was first used by Qureshi and co- w o r k e r ~ ' ~ , ~ ~ in determining the structure of monophosphoryl-lipid A from LPS (Salmonella typhimurium). The earliest lipid A study to use laser desorption mass spectrometry (LDMS) was reported by Seydel and LindnerI7 for lipid A from Rhodomicrobium vaniellii. A feature of LDMS (a forerunner of matrix- assisted laser desorption) is the high internal energy imparted to analyte species which causes specific frag- mentation of lipid A, enabling one to establish the location of fatty acids on the lipid A backbone.'8319 Plasma desorption mass spectrometry (PDMS) has also been employed for the examination of many kinds of lipid As from different bacterial families.20-22 Although PDMS studies of lipid A provide useful structural information, analyses suffer from the low mass resolution inherent to the linear time-of-flight mass spectrometers employed.

Electrospray ionization mass spectrometry (ESMS) is another ionization technique which has relatively recently distinguished itself by the ability to produce intact singly or multiply charged ions from proteins and other b i o p o l y m e r ~ . ~ ~ . ' ~ As first demonstrated in our

ESMS is particularly suited to lipid A structural studies. As perhaps the 'softest' of all mass spectrometric ionization techniques, under appropriate experimental conditions, electrospray ionization can yield minimal gas-phase fragmentations of target analytes (e.g. lipid A) to allow viewing of the profile of forms of lipid A produced as a result of hydrolysis of LPS.30 Additional information pertaining to lipid A structure can be obtained if collision-induced disso- ciation (CID) is employed to fragment ES-desorbed species either prior to the mass analyzer on a conven- tional mass spectrometer3' or, more elegantly, in the collision cell of a tandem mass ~pectrometer.~'

In general, to determine fully the lipid A structure of a certain bacterial strain, mass spectrometry must be used in conjunction with other analytical techniques. Fatty acyl groups may be hydrolyzed from lipid A by base or acid treatment, converted into the methyl ester forms and identified by gaslliquid chromatography

(GLC) or GLC/MS. Phosphate groups can be determined by chemical analysis or by 31P NMR, although exact fatty acyl group positions and anomeric configu- rations must be determined by two-dimensional nuclear magnetic resonance (2D-NMR).

Numerous studies have shown that, with certain exceptions, lipid A from many different bacterial sources share a common backbone structure: 8-1,6-linked D- glucosamine disaccharide (Fig. 1). Amide-linked fatty acyl side-chains from these lipid A species are usually found at the 2- and 2'-positions, while ester linkages of such side-chains appear at the 3- and 3'-positions, with phosphate groups at the l-and #-position of the disaccharide backbone. Additional ester-linked fatty acyl groups are also frequently found at the sites a or 8 to the carbonyl of both amide- and ester-linked side- chains. Differences between lipid As of the Salmonella minnesota and Escherichia coli type are found only in the number and kinds of fatty acyl groups. The fatty acyl group chain length can vary from C,, to c28.

Because of the frequently similar diglucosamine backbone structure, often unknown lipid A structures can be postulated using MS analysis alone. At present, the lipid As whose complete structures have been established are S. typhimurium G30/21, S. minnesota R595, E. coli and Rhodobacter sphaeroides ATCC 1 7023.3 '

A particular LPS that we have been investigating",' '9' 6,2 9 7 3' is that produced by Entero- bacter agglomerans, a Gram-negative bacterium commonly found in field cotton. LPS in cotton dust has been found to be the cause of respiratory diseases such as b y s s i n ~ s i s . ~ ~ The Pittsburgh animal test has shown that LPS from Enterobacter agglomerans can cause pulmonary function i m ~ a i r m e n t ~ ~ . As is true for other bacteria, the toxic principle of the LPS is believed to reside in the lipid A moiety. Previous work in our laboratory using both PDMS22 and ESMS3' has led to the assignment of the structure of lipid A from E . agglomerans shown in Fig. 1. It was determined that lipid A from E. agglomerans (EA) closely resembles the known structure of lipid A from S . minnesota (SM).37*38 Eight varieties of lipid A molecules differing only in the number of attached side-chains were present in the crude lipid A mixture obtained via acid hydrolysis of LPS (EA). It was not clear, however, whether these eight forms of lipid A existed initially as heterogeneous LPS varieties, or whether some forms were generated during the hydrolysis process required to liberate lipid A from the intact LPS.

Among many detoxification approaches which have been tried, acid and base treatments of E . agglomerans LPS have been proved to be effective at reducing the endotoxin level. 39 Base treatment4' is especially effective at reducing the pulmonary toxicity of cotton dust, as measured by changes in respiratory frequency and tidal volume in a guinea pig animal model s t ~ d y . ~ ~ , ~ ~ The mechanism of detoxification, however, has not been established. The purpose of this study was to determine the structural changes occurring to the lipid A portion induced by both acid and base treatments. These structural modifications will be related to literature reports of reduced endotoxin levels and reduced pulmonary toxicities resulting from chemical treatments. Longer term goals are to reveal the minimum structure

140 Y. WANG AND R. B. COLE

responsible for lipid A toxicity and to define the minimum treatment required to remove the toxicity of lipid A.

sity peaks in each mass spectrum originate from back- ground noise. The mass spectrometer was calibrated to give m/z values comprehending zero mass defect.

EXPERIMENTAL

Lipid A preparation

LPS produced by E . agglomerans ATCC 27996, (LPS (EA)) was the source of lipid A employed in this study. LPS was extracted from E . agglomerans ATCC 27996 cells by the hot water-phenol m e t h ~ d . ~ ' The method- ology for culturing the organism and isolating LPS has been described previously.22 To separate the lipid A portion from LPS, the following procedures were used: LPS (EA) was mixed with 1% acetic acid (in pyrogen- free water) to a concentration of 1-5 mg ml-' and then placed in an ultrasound bath for 1-2 min until the sample was fully dissolved. It was then heated at 100 "C in a water-bath for about 30 min, at which time a precipitate (lipid A) began to form. Heating was continued for another 5 min and afterwards the test-tube was removed from the water-bath and placed in an ice-bath. Precipitated lipid A was isolated by centrifugation (3000 rpm, 30 min, w4OC). The upper water layer was removed and the precipitate was washed with pyrogen- free water. The centrifugation and washing steps were repeated three times. Finally, the sample was lyophi- lized to give solid lipid A totalling 10-15% of the starting material weight. This preparation procedure was also employed to obtain lipid A from LPS of Salmonella minnesota (Re mutant strain R595) purchased from Sigma Chemical (St Louis, MO, USA).

Electrospray ionization mass spectrometry

Electrospray ionization (ES) mass spectra were acquired on a Vestec Model 201 single quadrupole mass spectrometer (PerSeptive Biosystems, Vestec, Houston, TX, USA) capable of analyzing ions up to m/z 2000. Direct infusion of sample solution was performed using a Sage syringe pump (Orion Research, Boston, MA, USA). Solutions containing 0.05-0.1 mg ml-' lipid A in methanol-chloroform (1 : 1) were infused into the ES chamber at a flow-rate of 2 p1 min-'. All mass spectrometric experiments were conducted in the negative-ion mode. The operating voltage of the ES capillary (stainless steel) was about -1.65 kV and the nozzle (counter electrode) voltage and the skimmer-collimator voltage difference were maintained at -290 V and -20 V, respectively. The temperature at the ES capillary tip was held at 55560°C. Each series of mass spectra was obtained under constant experimental conditions, over a total time interval of less than 1 h. Displayed mass spectra represent raw data obtained from the average of at least ten scans, each of 6 s duration. Post-acquisition data smoothing was not employed and the lowest inten-

RESULTS AND DISCUSSION

Hydrolysis of LPS

Lipid A is usually prepared by performing mild acid hydrolysis of LPS, which selectively hydrolyzes the KDO-glucosamine bond linking the core region and lipid A. Depending on the method used to extract LPS from bacteria, the LPS obtained may be in the form of S-type (with O-specific chain) or R-type (without 0- specific chain). R-type LPS, depending on how incom- plete the core structure may be, are further classified into Ra, Rb, Rc, Rd and Re types. Optimum conditions for hydrolysis of the various types of LPS to generate lipid A may differ substantially. Preparation conditions that can be varied include acid concentration, treatment time and temperature. Because the purpose of this study was to find out how lipid A structure becomes modified via chemical treatment with the goal of reducing and finally eliminating the toxicity, it was important that lipid A starting material be as intact as possible. In other words, lipid A should be released from the LPS in a gentle manner such that, ideally, initial acid hydrolysis breaks only the KDO-lipid A bond without disrupting other structural features including the fatty acyl chains on lipid A.

Various concentrations of acetic acid having pH values of 4.5, 3.9, 3.2 and 2.8 were used to perform the lipid A preparation from LPS (EA) in a water-bath at 100°C. At pH 4.5 and 3.9, even after fairly long treatment times (3 h), no lipid A precipitate was observed, indicating that these conditions were too mild. At pH 3.2, a small amount of lipid A precipitate appeared after 2 h of treatment, but the yield was very low. At pH 2.8 (1% HOAc), however, a precipitate began to form after 30-35 min. Additional precipitate formation continued for about 5 min, afterwhich the total lipid A (EA) product weighed about 13% of the starting material weight. LPS, of course, are amphipathic molecules com- posed of a hydrophilic sugar portion represented by the O-specific chain plus the core oligosaccharide and the hydrophobic lipid A portion (Scheme 1). The hydrophobic interactions of fatty acyl chains (including micelle formation at elevated concentrations) may diminish the accessibility of H,O+ to the KDO-lipid A bond. Higher acid concentrations (lower pH, e.g., pH 2.8) further shortened the hydrolysis time, but, as will be shown later, losses of the fatty acyl groups began to occur. For this reason, 1% acetic acid (pH 2.8) at 100 "C was chosen as a standard condition for lipid A preparation from LPS (EA). The same condition was used to generate lipid A from S . minnesota Re mutant strain R595 (SM). Compared with LPS (EA), the hydrolysis of LPS (SM) proceeded more quickly and in about 5-8 min the precipitation was complete; the product totalled 55% of the starting material weight.

HYDROLYSIS OF LIPID A AS MONITORED BY ESMS

Differences in the source LPS material for E . agglomerans and S . minnesota may explain the different lengths of time required for completion of lipid A formation and also the different yields produced. LPS (EA) is an R-type lipopolysaccharide containing a core oligosaccharide, although its complete structure has not yet been determined. LPS (SM), on the other hand, has a small number (two) of KDO groups linked with lipid A. Because of the hydrophobicity of the lipid A portion within LPS, LPS molecules tend to form micelles in aqueous solution. These micelles may impede hydronium ion attack at the KDO-lipid A linkage. Moreover, depending on the type of LPS, the micelle conformation can be different. This can lead to differences in accessibility of the hydronium ions to the KDO-lipid A bond. This may explain why mild acid hydrolysis (1% acetic acid, at 100 "C) proceeded relatively quickly for LPS (SM) (5-8 min) compared with LPS (EA) (30-35 min) to complete lipid A precipitation.

Lipid A from E . agglomerans (Fig. 1) was found in previous ~ o r k ~ ~ , ~ ~ to have a similar structure to that of lipid A from S. minnesota. Fully characterized heptaacyl structures identical with that shown in Fig. 1 have been r e ~ o r t e d ~ ~ , ~ ' for lipid A (SM). One report37 indicates the existence of heterogeneity at the R,' side-chain of lipid A (SM), similar to that shown in Fig. 1 for lipid A (EA). The lipid As from both E. agglomerans and S . minnesota used in our previous s t ~ d i e s ~ ~ * ~ O were obtained from the respective lipopolysaccharides using stronger acid conditions, higher temperatures and/or longer treatment times than those employed in this study. Previously, lipid A (EA) was obtained by 1% acetic acid treatment at 116 "C for 1 h; lipid A (SM) was obtained by 1.0 M HCl treatment at 100°C for 1 h. As was shown in ES mass spectra,30 at least eight varieties of lipid A existed in the crude preparation. The heterogeneity took the form of differing levels of acylation, i.e. different numbers (and in the case of R1', different types) of ester-linked hydrocarbon chains. At that time, it could not be determined whether the different levels of acylation existed within the intact LPS, or whether some less acylated varieties were formed via hydrolysis during the process of lipid A preparation from LPS.

To clarify this point, LPS from E . agglomerans and S . minnesota (R595) were prepared according to the above- mentioned (standard) mild conditions (1% acetic acid at 100°C). Once the formation of lipid A was apparently complete, the reaction vessel was placed in ice-water. Immediately afterwards, the lipid A precipitate was isolated by centrifugation and lyophilization. Figures 2(a) and 3(a) show the ES mass spectra of lipid A from E. agglomerans and S. minnesota respectively, obtained under the mild conditions described above. In comparing Figs 2(a) and 3(a), many features are similar for the mass spectra of the two lipid A types. In both spectra, m/z 1956 is the base peak, with a second major peak at m/z 1717. Based upon the known structure of lipid A from S . minnesota we know that these peaks correspond to the two major forms which are heptaacyl and hexaacyl-lipid A, respectively. In contrasting these mass spectra with those of comparable lipid As produced under harsher condition^,^^ it is clear that substantial acyl side-chain hydrolysis occurred during the previously employed lipid A preparation procedure. We

a 5

c

LA Ilcn

141

4 ?ua

IUNI 12m1 14W I NKI I an1 21m OM2

l c

Figure 2. Negative-ion ES mass spectra of lipid A (EA) after mild acid treatment (1% acetic acid, 100°C): (a) 0 ; (b) 30; (c) 60; (d) 120 min.

142 Y.

1

WANG AND R. B. COLE

I ( Y X I 12IXI 11111t IhlXl IUXI ?IXXI d l

Figure 3. Negative-ion ES mass spectra of lipid A (SM) after mild acid treatment (1% acetic acid, l0O'C): (a) 0; (b) 10; (c) 20; (d) 30 min.

consider the current preparation conditions to be causing a minimum of acid hydrolysis. The mass spectral peaks appearing in both Figs 2(a) and 3(a) are interpreted in Table 1 (other major mass spectral peaks which appear in later spectra are also assigned in Table 1).

From these two mass spectra and the interpretation in Table 1, the resemblance of the two lipid As from LPS (EA) and LPS (SM) is clear, in agreement with our previous report.30. Slight differences existed between the two ES mass spectra such as the appearance of m/z 1972 [Fig. 2(a)] which has been e ~ p l a i n e d ~ ~ * ~ ~ by the heterogeneity at R,' (Fig. 1) and other slight differences which are likely to be attributable to further heterogeneity of LPS. In comparing the mass spectra, it is evident that the lipid A (EA), whose mass spectrum is shown in Fig. 2(a), has undergone a higher level of hydrolysis than the lipid A (SM), whose mass spectrum is shown in Fig. 3(a). This is evidenced by the fact that the relative peak intensities at m/z 1745, 1507 and 1491, against m/z 1956 shown in Fig. 2(a), are higher than the corresponding intensity ratios in Fig. 3(a). The additional hydrolysis may be attributable to the longer treatment time employed during lipid A (EA) preparation from LPS as compared with lipid A (SM). Fur- thermore, the ES mass spectrum of lipid A (EA) is observed to contain additional peaks at m/z 1797, 1587 and 1571 [Fig. 2(a)], which correspond to diphosphoryl-lipid A species (Table 1) which are absent from the lipid A (SM) spectrum [Fig. 3(a)]. The use of the minimum hydrolysis condition to generate lipid A demonstrates that lipid As from E . aggZomerans and S . minnesota exist mainly in heptaacyl and hexaacyl forms, although EA lipid A has been shown3' to consist of two distinct heptaacyl along with two hexaacyl varieties. Other forms containing fewer acyl side-chains observed previously30 are thus largely produced during the process of LPS hydrolysis.

Influence of treatment time on acid hydrolysis

Among the factors which can be adjusted to control the hydrolysis process are acidity, temperature and treatment time. The influence of treatment time was investi- gated first. Lipid A from each bacterium was treated separately in 1% acetic acid at 100°C. Time zero indicates the time at which the mild acid hydrolysis of LPS, i.e. lipid A preparation, was just complete under minimum hydrolysis conditions and the intact lipid A had fully precipitated. Three other identical samples were run in parallel for longer treatment times (30, 60 and 120 min for EA and 10,20 and 30 min for SM, after the lipid A preparation was complete). After the proper time interval had elapsed, treatment was stopped by placing the test-tube in an ice-bath, followed immediately by washing, centrifugation and lyophilization; the reaction products were analyzed by ESMS.

Figures 2(a-d) and 3(a d) show the ES mass spectra of lipid A (EA) and lipid A ( S . minnesota R595) respectively, corresponding to the different acid hydrolysis times. In each figure, a clear, progressive decrease in the relative intensity of the m/z 1956 peak is observed with a concomitant increase in that of m/z 1717. From the


Table 1. Interpretation of negative-ion ES mass spectra of lipid A from E. ag~lumerlp ans and S. minnesotaa

rn II

1972 1956 1928 1813b 1 797b 1745 1733 1717 1689 1587b 1571 1507 1491 1361 1281 11 78b 1054 1036

Empirical lormula

OH

R 4

Lipid A backbone with different acvl-linked side-chains

A , R, A 3 R,

HN-OM' L-OM' HM' P-OM' M-OM' L-OM' HM' P-OM'

Less C2H, relative to miz 1956 HN-OM' L-OM' HM' H M' M-OM' L-OM' HM' H M' H N' L-OM' HM' P-OM' HN-OM' L-OM' HM' ti M' M-OM' L-OM' HM' H M'

HN' L-OM' HM' H M' M-OM' L-OM' H' H M' HN' L-OM' HM' H M' M-OM' L-OM' H' HM' H N ' C L-OM' H' H M' HN'C L-OM' H ' H M' HN'C H M' H' HM' H' L-OM' H' HM'

Less C2H, relative to m/z 171 7

Less H,O relative to miz 1054

'P= palmitoyl (-C16), M = myristoyl (-C,,), H M = hydroxymyristoyl (-C,,OH), L = lauroyl ( -C12), OM = oxymyristoyl (-C,,O), H -hydrogen Ions representative of diphosphoryl lipid A , the second phosphoryl group (not shown)

is located in the place of the ( 1 -position) axial hydroxyl group on the extreme right of the structure

HM' could also be located on R, instead of A,

structure of lipid A in Fig. 1 and the mass spectral interpretation in Table 1, the m/z 1956 peak is known to possess an additional palmitoyl group R,' relative to m/z 1717. The gradual change in the relative intensities of the m/z 1956 and 1717 peaks indicates that during the hydrolysis process, the palmitoyl group of heptaacyl-lipid A is hydrolyzed, thereby forming hexaacyl-lipid A.

The rate equation for acid hydrolysis of an ester may be expressed as u = k[ester][H,O+]. At pH 2.8, [H,O+] = 1.6 x lo-, M, while [ester] = 1.4 x M (heptaacyl-lipid A) at time zero in our (EA) experiments. The initial heptaacyl-lipid A concentration was esti- mated from the intensity of the m/z 1956 peak relative to all other lipid A peaks shown in Fig. 2(a) (i.e. - 20%), considering a total lipid A (EA) product weight of 13% of the LPS (EA) starting material weight which was initially present at 1.0 mg m1-l. With H,O+ present in large excess relative to lipid A, the reaction kinetics can be simplified to pseudo-first order (u = k'[heptaacyl- lipid A]). At time zero, the decomposition of LPS to produce both heptaacyl- and hexaacyl-lipid As was complete as verified in separate experiments by removing the lipid A precipitate by centrifugation, followed by restarting the 100°C water-bath treatment of the remaining solution. No additional precipitate was found during a further 2 h of treatment. It is therefore reason-

able to assume that during further hydrolysis of lipid A, any increase in the peak intensity of hexaacyl-lipid A was due solely to hydrolysis of the palmitoyl group from heptaacyl-lipid A, and that this loss of the palmi- toy1 group represents the first step of heptaacyl-lipid A decomposition while other products of heptaacyl-lipid A decomposition are negligible. It is also reasonable to presume that the difference in the electrospray ionization efficiencies of hexaacyl-and heptaacyl-lipid As is negligible. Lastly, we assume that over the course of acid treatment, the hexaacyl-lipid A (m/z 1717) under- goes negligible further hydrolysis, which is justifiable by the observation that lower mass peaks (e.g. m/z 1507 and 1491) remain low-intensity peaks under all conditions shown in Figs 2 and 3.

With these assumptions, the relative peak intensities of m/z 1956 and 1717 should reflect the relative concentrations of the heptaacyl and hexaacyl forms of lipid A, respectively, in the crude mixture. Defining C, and C, as the heptaacyl-lipid A concentration at time zero and time t respectively, a plot of In(C,/C,) us. time shows a linear relationship in accordance with the presumed pseudo-first-order reaction [Fig. 4(a)]. From the slope of the curve, k' [the pseudo-first-order rate constant for heptaacyl-lipid A hydrolysis to form hexaacyl-lipid A (EA)] is 3.3 x min-' at 100°C. An analogous experiment was performed using lipid A from S. minne-


-0.45 1 , . . , I I ’ ‘ I ’

0 20 40 60 80 100 120 time (min)

-o.o:h”; k = -8.4~10” (min-’)

I ” 0 5 10 15 20 25 30

time (min)

Figure 4. Plots of In(C,/C,) vs. time (rnin) for (a) lipid A (EA) and (b) lipid A (SM) during further acid hydrolysis.

sota. The ES mass spectra of lipid A (SM) as a function of treatment time are shown in Fig. 3(a4j . Similarly, a plot of In(C,/C,) us. time yields a linear relationship as expected for a pseudo-first-order reaction [Fig. 4(b)]. The slope of the curve (k’) reveals the pseudo-first-order rate constant for palmitoyl ester chain hydrolysis to be 8.4 x lo-, min-’ at 100 “C for lipid A (SM).

The difference in the k‘ values for lipid A (EA) and lipid A (SM) may be partially attributable to the existence of diphosphoryl-lipid A components (Table 1) in lipid A (EA) which can undergo competitive hydrolysis processes. In addition, the phosphate group at position 1 is known to be acid labile. It seems, however, that the palmitoyl group is more acid labile than the phosphate group at position 1, because the relative intensity of the peak at m/z 1797 (diphosphorylhexaacyl-lipid A, Fig. 2) did not undergo much change under the mild acid treatment conditions. As will be explored in the next section, when more concentrated acid was used the phosphoryl group at position 1 was hydrolyzed so that diphosphoryl species were not observed.

Influence of acid concentration

The influence of acid concentration on the hydrolysis process was studied by using acetic acid at different concentrations (i.e. 2%, 5% and 10% solutions having pH values of 2.7, 2.5 and 2.4, respectively) to perform hydrolysis of LPS (EA). In the three cases, it took about 28, 25 and 20 min, respectively, for lipid A to precipitate. The ensuing lipid A work up was the same as that described previously. ES mass spectra of the obtained products are shown in Fig. 5(a-c). It is evident that higher acid concentrations increasingly hydrolyze the ester-linked side-chains during the process of lipid A preparation when these spectra of lipid A acquired after treatment with increasing acetic acid concentrations [Fig. 5(a-c)] are compared with that in Fig. 2(a) which was acquired after treatment under the mildest (1% acetic acid) condition. The sequential decrease in the relative intensity of the m/z 1956 (parent heptaacyl) peak observed when comparing Figs 2(a) and 5(aj, (b) and (c) (order of increasing treatment acidity) and the dominance of the m / z 1717 (hexaacyl) peak are consis- tent with results given in the influence of treatment time section, i.e. the palmitoyl group is the side-chain most labile to acid hydrolysis. All the mass spectral peaks with lower m / z (e.g. 1507, 1491, 1281 and 1054) can be interpreted (Table 1) as losses of various ester-linked side-chains from the parent heptaacyl (after first losing a palmitoyl group) or from hexaacyl-lipid A.

Other than the heptaacyl parent at m/z 1956, the peak at m/z 1745 (arising from removal of R,’ from the heptaacyl parent) represents the only ion to contain the palmitoyl group at R,‘ (see Table 1). This implies that the removal of R,’ from the parent is the only decomposition which is competitive with palmitoyl hydrolysis. This provides evidence that the hydrolysis of R1’ occurs with the second fastest rate constant (after the R,‘ palmitoyl group). Based on the acid hydrolysis mechanism, in order to initiate hydrolysis of an ester-linked side- chain, the carbonyl oxygen needs to be accessible to H,O+. From the lipid A structure (Fig. l), it seems reasonable that the R,’ and R,‘ ester chains are the least sterically hindered, which can explain why they are the most susceptible to H 3 0 + attack and subsequent hydrolysis. Starting from a parent heptaacyl-lipid A (which has already undergone an initial palmitoyl loss) or from hexaacyl-lipid A, the m / z 1491 ion can only be formed by decomposition of the R, side-chain (Table 1). On the other hand, from the same starting point, hydrolysis of the R, side-chain would produce an ion at m/z 1281. Because the peak at mjz 1491 appears as a hydrolysis product prior to the appearance of m/z 1281 [Fig. 2(a)], it may be deduced that the R, hydroxymyristoyl group at position 3 is more labile than the R, side-chain at position 3’. The R, hydroxymyristoyl chain must also be more labile than the lauroyl chain at position 2‘ because the latter is present in every mass spectrum displayed in Figs 2 and 5, i.e. it was never removed by hydrolysis. The 3’-oxymyristoyl group is also relatively resistant to hydrolysis, but the presence of the ion at m/z 1054 [Fig. 5(c)] containing only amide-linked side- chains and the lauroyl group at position 2 (Table 1) suggests that the 3’-oxymyristoyl group is more labile than the lauroyl moiety. From the above observations,


i a ‘1 s

C 1

IWO I zoo 1400 1600 I800 zoo0 d 2

Figure 5. Negative-ion ES mass spectra of lipid A (EA) obtained for LPS hydrolysis using acetic acid of different concentrations: (a) 2%; (b) 5%; (c) 10%.

an approximate order of hydrolysis of the ester-linked Theoretically, the hydrolysis process of each individ- side-chains during further acid hydrolysis was deduced: ual ester chain should obey pseudo-first-order reaction R,‘ (palmitoyl) > R,’ (myristoyl or hydroxy- kinetics as demonstrated for the palmitoyl group. myristoyl) > R, (hydroxymyristoyl at position 3) > R, However, when more concentrated acid was used, (oxymyristoyl group at position 3’) > R1‘ (lauroyl). several hydrolysis processes proceeded simultaneously,


complicating the overall reaction, and the disap- pearance of the parent structure could no longer be described by simple kinetics. It is also useful to note that the m/z 1797 peak corresponding to diphosphoryl- lipid A (EA) which has lost the palmitoyl group at R,' (Table 1) is the most prevalent diphosphoryl peak appearing in Fig. 2(a-d). The observation of this large m/z 1797 peak provides evidence that, even in the presence of a second phosphoryl group at the 1-position (not far from the R,' palmitoyl group, Fig. I), the R,' palmitoyl group is still the acyl side-chain which is most readily hydrolyzed. This is important to note because the native LPS (EA) contains a phosphoryl group at the 1-position. This evidence suggests that the above hydrolysis order would remain the same in the native LPS.

Effect of base hydrolysis

Domelsmith and ~ o - w o r k e r s ~ ~ * ~ ~ found that LPS from EA treated with 0.05 M NaOH in 95% EtOH was most effective at reducing the pulmonary toxicity compared with other solvent systems (e.g. HC1, H,P04, K2C03 , dimethyl sulfoxide, dimethylformamide and diethylene glycol, each in the same 95% EtOH solvent). The reason that non-aqueous solutions were employed is that water-washed cotton is difficult to use in textile processing because waxes and inorganic salts have been removed from the cotton fiber. Inorganic salts, left largely intact after ethanol washing, are believed to improve the ability to process cotton fibers by dissi- pation of static Because lipid A is the moiety believed to be responsible for the toxicity of LPS, lipid A 'starting material' was first obtained from LPS (EA) using mild acid hydrolysis as described above.

Base treatment of lipid A was initiated by loading a 5 ml vial with about 0.5 mg of the lipid A preparation. The sample was first dissolved in 500 pl of CHC1, , after which 500 pl of 0.05 M NaOH in 95% EtOH was added. The solution was then placed in an ultrasonic bath for 1-2 min. Four vials were heated on the same heating block at 65 "C for different lengths of time, i.e. 15, 30,45 and 60 min. After the treatment time had expired, each vial was cooled and neutralized by addition of 500 p1 of 1% acetic acid. The post-treatment products were extracted with 1.0 ml of chloroform. Emulsions which formed during extraction were broken by centrifugation (3000 rpm). After three extractions with chloroform, the organic layer was removed via lyophilization. The lyophlized product was dissolved in 1.0 ml of methanol-chloroform (1: 1) and was now ready for ESMS analysis.

ES mass spectra of base-treated lipid A are shown in Fig. 6, beginning with the spectrum of the lipid A starting material [Fig. 6(a)] obtained by mild acid hydrolysis of LPS (EA) with no base treatment, defined as time zero. The ES mass spectra of the products of 15, 30 and 45 min base treatment are shown in Fig. 6(b), (c) and (d), respectively; they clearly reflect changes attributable to progressive base hydrolysis. Ions representative of products which have lost ester-linked side-chains via hydrolysis from the parent monophosphoryl heptaacyl- or hexaacyl-lipid A (EA) appear at m/z 1507, 1491, 1281 and 1054. Assignments for these peaks are included in

.. ..

1111 11111 n ln l 14111 IMX)

1

nd,.

HII IWII IIIM IMI I I ~ i ~ u i nnxi n h

Figure 6. Negative-ion ES mass spectra of lipid A (EA) after base treatment (0.05 M NaOH in 95% ethanol, 65°C): (a ) 0; (b) 15; (c) 30; (d) 45 min.

Table 1. From Fig. 6(a), we know that the lipid A starting material contains some diphosphorylhexaacyl-lipid A (m/z 1797). The base treatment series shows that the diphosphorylhexaacyl-lipid A was resistant to decom-


position [e.g. see Fig. 6(b-d)], but was eventually com- pletely hydrolyzed after 60 min of treatment (not shown).

As interpreted in Table 1, the peaks at m/z 1587, 1571, 1361 and 1178 were formed by hydrolysis of various ester side-chains from parent diphosphoryl-lipid A. Added Coulombic repulsion originating from the presence of the second ionizable phosphoryl group may inhibit, to some degree, approach of hydroxide ions to the diphosphoryl-lipid A, thus slowing the hydrolysis process relative to monophosphoryl-lipid A. Attack of the hydroxide ion at the carbonyl carbon of the ester chains is the initial step in the base hydrolysis process. Under the base treatment conditions employed, hydrolysis of ester side-chains proceeded much faster and almost simultaneously as compared with acid treatment conditions. For this reason, a kinetic model describing individual ester chain base hydrolysis could not be established.

A noteworthy feature of ES mass spectra obtained after base treatment of lipid A (EA) is the appearance of peaks [e.g. m/z 875 and 857 in Fig. 6(c) and (d)] which cannot be rationalized as acyl chain losses from intact lipid A. These peaks took on increased prominence in the mass spectrum as base treatment progressed. When the treatment time reached 1 h, no peaks above m/z 1000 remained; the lower mass portion of the ES mass spectrum corresponding to 60 min base tratment (obtained later, and under higher sensitivity conditions than existed during the acquisition of Fig. 6) appears in Fig. 7. Ions at m/z 199, 227, 243 and 255 (Fig. 7) represent L, M, HM and P, respectively, in their carboxylate anion forms which were released from lipid A (EA) during the hydrolysis process. Remaining prominent peaks at m/z 937, 875, 857, 477 and 468 are likely all to be structurally related to the m/z 955 peak. Relative to m/z 955, they may be formed by removal of H,O (m/z 937), or by removal of a proton from singly charged m/z 955 or 937 to form doubly charged m/z 477 or 468, respectively. Moreover, replacement of a phosphoryl group by a hydroxyl group on m/z 955 or 937 will form

m/z 875 or 857, respectively. While the relationship between the above six peaks seems well founded, the identity of the diphosphoryl m/z 955 is less apparent. We propose the structure shown inset in Fig. 7 to represent m/z 955. The formation of this species, however, must involve an unusual reductive pyranose ring opening for each glucosamine moiety. The mechanism involved in such a reduction is not obvious and electro- chemical processes occurring at the negatively charged ES needle cannot be ruled out. When stronger base (>0.1 M NaOH) or an even longer treatment time (> 1 h) was employed, it appears that even the remaining phosphoryl group was hydrolyzed because no negatively charged lipid A fragments other than the carboxylate anions could be detected by the ES mass spectrometer .

CONCLUSION

ESMS has proved to be an efficient technique for lipid A structural studies. ES mass spectra of lipid A from E . agglomerans obtained via mild acid hydrolysis revealed that lipid A was produced mainly in two forms, heptaacyl and hexaacyl, implying that a large amount of the less acylated lipid A products obtained under harsher conditions3’ were formed during the procedure used to prepare lipid A from LPS. When acid hydrolysis of lipid A was performed, ester-linked side-chains were progressively removed. Among these ester side-chains, the palmitoyl group was found to be the most readily hydrolyzed; the decomposition conformed to pseudo- first-order reaction kinetics. The approximate order of acid hydrolysis of the ester side-chains was determined to be R,’ (palmitoyl) > R,’ (myristoyl or hydroxymyristoyl) > R3 (myristoyl group at position 3) > R, (oxymyristoyl group at position 3’) > R,’ (lauroyl). Under the conditions employed, the use of base hydrolysis was even more effective at removing

1 OH

100 500 lob0 m/z

Figure 7. Lower mass region of the negative-ion ES mass spectrum of lipid A (EA) after 60 min of base treatment. Inset: proposed structure of the m/z 955 ion (the charge may be located on either phosphoryl group).

148 Y. WANG A N D R. B. COLE

acyl side-chains. The exact number and location of ester chains which need to be removed to eliminate the toxicity remain to be determined, as does the minimum structure of lipid A responsible for toxicity.

Use of sodium hydroxide (0.05 M in 95% ethanol) to treat LPS (EA) was consistently effective in lowering the endotoxin level, as measured by the Limulus amebocyte lysate (LAL) assay, and also effective in reducing the pulmonary toxicity of cotton dust, as monitored by changes in respiratory frequency and tidal volume in a guinea pig animal mode1.40,42v43 Although acid treatment could also reduce the endotoxin level of LPS (EA), it was not so effective at removing the pulmonary toxicity. Based on this study's deduced structural changes to lipid A, evidence has been provided that the decrease in endotoxin level after either acid or base treatment may be attributed largely to the removal of acyl side- chains. Furthermore, there may also exist a mechanism

for eliminating the pulmonary toxicity from LPS (EA) which goes beyond hydrolysis of the acyl side-chains. The possibilities brought forth that the pyranose rings of the lipid A disaccharide backbone may be opened, or the phosphoryl groups removed, during base treatment are conceivable explanations.

Acknowledgements

Partial financial support for this research was provided by the Loui- siana Education Quality Support Fund through grant No. (1991- 1994)-RD-A-36. Financial support was also received from the USDA through cooperative agreement No. 58-6435-4-104. We thank Dr A. J. DeLucca of SRRC, USDA, for supplying the LPS from E . agglomerans. We also thank Drs Linda Domelsmith, Mary-Alice Rousselle and William Franklin of SRRC, USDA, for helpful discussions and valu- able technical support.

REFERENCES

1 .

2. 3.

4.

5.

6.

7.

8

9

10

11

12 13

14

15

16

17

18

19

20.

21. 22.

23.

24.

C. Galanos, J. Roppel, J. Weckesser. E. T. Rietschel and H. Mayer, Infect. lmmun. 16, 407 (1 977). E. T. Rietschel and H. Brade, Sci. Am. August, 54 (1 992). N. Qureshi, K. Takayama, K. C. Meyer, T. N. Kirkland, C. A. Bush, L. Chen, R. Wang and R. J. Cotter, J. Biol. Chem. 266, 6532 (1991). S. Basu, J. Radziejewska-Lebrecht and H. Mayer, Arch. Microbiol. 144, 21 3 (1 986). R. Bhat, A. Marx. C. Galanos and R. S. Conrad, J. Bacteriol. 172,6631 (1 990). 0. Luderitz, M . A. Freudenberg, C. Galanos, V. Lehmann, E. T. Rietschel and 0. H. Shaw, in Microbial Membrane Lipids, edited by S . Razin and S. Rottem. Vol. 17, p. 79. Academic Press, New York (1 982). J. Y. Homma and K. Tanarnoto, in Bacterial Endotoxins: Chemical Biological and Clinical Aspects, edited by J. Y. Homma, S. Kanegasaki, 0. Luderitz, T. Shiba and 0. West- phal. p. 159. Verlag Chemie, Weinheim (1 984). T. Shiba and S. Kusumoto, in Handbook of Endofoxins, Vol. 7: Chemistry of Endotoxins, edited by E. T. Rietschel, p. 284. Elsevier, Amsterdam (1 984). T. Yasuda, S. Kanegasaki, T. Tsurnita, T. Tadakuma, J. Y. Hornma, M. Inage, S. Kusurnoto and T. Shiba, T., Eur. J. Biochem. 124,405 (1 982). H. Takada, S. Kotani, S. Tanaka, T. Ogawa, I. Takahashi, M. Tsujimoto. T. Komuro, T. Shiba. S. Kusumoto, e t a / . , fur. J. Biochem. 175, 573 (1 988). P. V. Salimath, J . Weckesser, W. Strittmatter and H. Mayer, Eur. J. Biochem. 136, 195 (1 983). M. Romano and J. Hawiger, J. Biol. Chem. 265,1765 (1990). M. Caroff, C. Deprun, D. Karibian and L. Szab0.J. Biol. Chem. 266,18543 (1 991 ). C. M. John, J. McLeod Griffiss, M. A. Apicella, R. E. Mandrel1 and 8. W. Gibson, J. Biol. Chem. 266,19303 (1 991 ). N. Qureshi, K. Takayama and E. Ribi, J. Biol. Chem. 257, 1 1 808 (1 982). N. Qureshi, K. Takayama, D. Heller and C. Fenselau J. Biol. Chem. 258,12947 (1 983). U. Seydel and B. Lindner. Springer Ser. Chem. Phys. 25, 240 (1 983). U. Seydel, B. Lindner, H.-W. Wollenweber and E. T. Rietschel, Eur. J. Biochern. 145, 505 (1 984). R . J. Cotter, J. P. Honovich, N. Qureshi and K. Takayama, Biomed. Environ. Mass Spectrom. 14, 591 (1 987). D. Karibian, C. Deprun, L. Szabo, Y. LeBeyec and M. Caroff, lnt J. Mass Spectrom. Ion Processes 11 1, 273 (1 991 ). R . J. Cotter, Anal. Chem. 60, 781A (1 988). R . B. Cole, L. N. Dornelsmith, C. M. David, R. A. Laine and A. J. DeLucca, Rapid Commun. Mass Spectrom. 6, 61 6 (1 992). J. B. Fenn, M. Mann, C. K. Meng, S. F. Wong and C. M. Whitehouse. Mass Spectrom. Rev. 9. 37 (1 990). E. C. Huang, T. Wachs, J. J. Conboy and J. D. Henion, Anal.

25.

26.

27.

28.

29.

30.

31. 32.

33.

34.

35.

36.

37.

38

39.

40

41

Chem. 62, 713A (1 990). A. K. Harrata. L. N. Domelsmith and R. B. Cole, in Book of Abstracts, 12th lnternational Mass Spectrometry Conference (Amsterdam, 26-30 August 1991 ), p. 287 (1 991 ). R. B. Cole, A. K. Harrata and L. N. Domelsmith, in Proceedings of the 40th ASMS Conference on Mass Spectrometry and Allied Topics (Washington, DC. 31 May-5 June 1992), p. 1647 ( 1 992). R . B. Cole and A. K. Harrata, Rapid Commun. Mass Spectrom. 6, 536 (1 992). R . B. Cole and A. K. Harrata, J. Am. SOC. Mass Spectrom. 4, 546 (1 993). A. K. Harrata, L. N. Domelsmith and R. B. Cole, in Cotton Dust Proceedings-1 7th Cotton Dust Research Conference. edited by R. R . Jacobs, P. J. Wakelyn and L. N. Domelsmith), p. 310. National Cotton Council, Memphis, TN (1 993). A. K. Harrata, L. N. Domelsmith and R. B. Cole, Biol. Mass Spectrom. 22, 59 (1 993). S. Chan and V. N. Reinhold.Ana1. Biochem. 218, 63 (1 994). E. T. Rietschel, L. Brade, B. Lindner and U. Zahringer, in Bac- terial Endotoxic Lipopolysaccharjdes, edited by D. C. Morrison and J. L. Ryan, Vol. 1. p. 3. CRC Press, Boca Raton. FL (1 992). D. Honeybourne, D. S. Wales, A. Watson, W. R . Lee and B. F. Sager, Byssinosis-Causative Agent and Clinical Aspects. Shirley Institute Publication S.43, Shirley Institute, England (1 982). M. A. Ellakkani, Y. C. Alarie, D. A. Weyel, S. Mazumdar and M. H. Karol, Toxicol. Appl. Pharmacol. 74, 267 (1 984). L. K. Ryan and M. H. Karol, Am. Rev. Respir. Dis. 140. 1429 (1 989). 0. Griffiths-Johnson, L. K. Ryan, K. Spear and M. H. Karol, in Cotton Dust Proceedings--1 3th Cotton Dust Research Con- ference, edited by R. R. Jacobs and P. J. Wakelyn, p. 101. National Cotton Council, Memphis, TN (1 989). E. T. Rietschel, H.-W. Wollenweber, H. Brade, U. Zahringer, B. Lindner, U. Seydel. H. Bradaczek, G. Barnickel, H. Labis- chinski and P. Giesbrecht, in Handbook of Endofoxins, Vol. 1: Chemistry of Endotoxins. edited by E.T. Rietschel, p. 187. Elsevier, Amsterdam ( 1 984). N. Qureshi, P. Mascagni, E. Ribi and K. Takayama, J. Biol. Chem. 260, 5271 (1 985). L. N. Domelsmith and M. A. Rousselle, in Cotton Dust Proceedings-I Ith Cotton Dust Research Conference, edited by R. R. Jacobs and P. J. Wakelyn, p. 105. National Cotton Council, Memphis, TN (1987). M. H. Karol, A. Ogundiran, C. Gatty, P. Millner, L. N. Domels- mith and M. A. Rousselle, in Cotton Dust Proceedings-1 7th Cotton Dust Research Conference, editerd by R. R. Jacobs and P. J. Wakelyn, p. 116. National Cotton Council, Memphis, TN (1987). C. Galanos, 0. Luderitz and 0. Westphal, Eur. J. Biochem. 9,


245 (1969). edited by R . R. Jacobs, P. J. Wakelyn and L. N. Domelsmith,

Proceedings-13th Cotton Dust Research Conference, edited 44. L. N. Domelsmith, M. A. Rousselle, in Cotton Dust by R. R. Jacobs and P. J. Wakelyn. p. 97. National Cotton Proceedings-72th Cotton Dust Research Conference, edited Council, Memphis, TN (1989). by R. R. Jacobs and P. J. Wakelyn, p. 76. National Cotton

43. L. N. Domelsrnith, A. J. DeLucca and J. J. Fischer, in Cotton Council, Memphis, TN (1988). Dust Proceedings-1 4th Cotton Dust Research Conference,

42. L. N. Dornelsmith and J. J. Fischer, in Cotton Dust p. 13. National Cotton Council, Memphis, TN (1 990).

Acid and base hydrolysis of lipid A fromEnterobacter agglomerans as monitored by electrospray...

Documents

Transcript of Acid and base hydrolysis of lipid A fromEnterobacter agglomerans as monitored by electrospray...