© 2009 Robert Tucker - University of Florida

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DESIGN, FABRICATION, AND OPERATION OF HYBRID BIONANODEVICES FOR BIOMEDICAL APPLICATIONS

By

ROBERT MATTHEW TUCKER

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2009

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© 2009 Robert Tucker

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To my Family and Friends, for their unwavering support

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ACKNOWLEDGMENTS

First and foremost, I thank my advisor Dr. Henry Hess for his guidance, friendship, and

support throughout the past seven years. It is difficult to imagine where I would be without his

mentorship, but I would undoubtedly be poorer without it.

I thank my committee members, Dr. Christopher Batich, Dr. Richard Dickinson, Dr.

Laurie Gower, and Dr. Scott Perry for their constructive criticism, advice, and teachings. I thank

Dr. Viola Vogel for her support during my initial foray into the research world as a scared

undergraduate. I thank all of the Hess Group members, past and present, without whom it

certainly would not have been much fun: Ashutosh Agarwal, John Clemmens, Bob Doot, Isaac

Finger, Dr. Thorsten Fischer, In-kook Jun, Parag Katira, Greg Lee, Isaac Luria, Elizabeth

Mobley, Dr. Takahiro Nitta, Scott Phillips, Sujatha Ramachandran, Jason Rudman, Ajoy Saha,

Shruti Seshadri, and Yoli Smith.

I thank Dr. Stefan Diez for the opportunity to study in Dresden, Germany at the Max

Planck Institute for Molecular Cell Biology and Genetics. Everyone at the institute aided me in

some way, but in particular I thank the members of the Diez Group: Michael Berndt, Corina

Bräuer, Gero Fink, Veikko Geyer, Dr. Leonid Ionov, Till Korten, Dr. Cecile Leduc, Doreen

Naumburger, Bert Nitzsche, Cordula Reuther, and Felix Ruhnow. I also thank the Fulbright

Program for the opportunity to live and study in Germany, a truly amazing experience.

I extend a heartfelt thanks to Karen, Dustin, and Paige Hegland and Keith and Pauline

Leavitt for their support during my time in Gainesville. I thank Jamie Phifer for her friendship

and answering my never-ending medical questions. I thank Sara Totten for her computer and

canine-training expertise. As always, I thank my Seattlites for inspiring me. More than words can

ever express, I thank my parents, brother, and sister for their support in everything I do. Finally, I

thank my dog Boltzmann, who kindly chose to not eat my laptop, despite eating everything else.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS.................................................................................................................... 4

LIST OF FIGURES .............................................................................................................................. 7

ABSTRACT .......................................................................................................................................... 9

CHAPTER

1 INTRODUCTION....................................................................................................................... 11

Engineering at the Cellular Level............................................................................................... 12 Hybrid Bionanodevices............................................................................................................... 16

2 BACKGROUND AND EXPERIMENTAL TECHNIQUES .................................................. 19

The Cytoskeleton......................................................................................................................... 19 Intracellular Transport ................................................................................................................ 20 In-vitro Motility Assay ............................................................................................................... 21 Wide-field Epi-fluorescence Microscopy .................................................................................. 22 Confocal Microscopy .................................................................................................................. 24

3 FABRICATION: DIRECT-WRITE, RAPID-PROTOTYPING OF BIOFUNCTIONAL PROTEINS ON THERMORESPONSIVE SURFACES ......................................................... 32

Introduction ................................................................................................................................. 32 Preparation of PNIPAM Surfaces .............................................................................................. 33 LHC-based Photopatterning of Proteins .................................................................................... 34 Confirmation of the LHC Effect in Gliding Motility Assays ................................................... 36 Testing the Functionality of Patterned Proteins ........................................................................ 37

4 STABILIZATION: CHARACTERIZATION OF PROTEIN ACTIVITY FOR THE DEVELOPMENT OF TEMPERATURE-INSENSITIVE HYBRID DEVICES ................... 42

Introduction ................................................................................................................................. 42 Results .......................................................................................................................................... 44 Discussion .................................................................................................................................... 44 Materials and Methods ................................................................................................................ 46

Kinesin and Microtubule Preparation ................................................................................. 46 Variable Temperature Motility Assay ................................................................................ 46 Measurement of Temperature ............................................................................................. 47 Measurement of Velocity .................................................................................................... 47

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5 CONTROL: ADAPTING CELLULAR CONTROL STRATEGIES TO HYBRID BIONANODEVICES ................................................................................................................. 50


Kinesin and Microtubules ................................................................................................... 58 Caged-ATP and Hexokinase ............................................................................................... 58 Motility Assays and Microscopy ........................................................................................ 58 Velocity and Radius Measurements ................................................................................... 60 Numerical Model ................................................................................................................. 60 Analytical Model ................................................................................................................. 62

6 ACTUATION: ASSEMBLY OF AN ISOPOLAR MICROTUBULE ARRAY TO PRODUCE MACROSCALE FORCES USING NANOSCALE COMPONENTS ............... 67


Kinesin and Microtubules ................................................................................................... 71 Microtubule Array Preparation ........................................................................................... 71 Imaging and Measurement .................................................................................................. 72

7 CONCLUSION AND OUTLOOK ............................................................................................ 76

LIST OF REFERENCES ................................................................................................................... 78

BIOGRAPHICAL SKETCH ............................................................................................................. 91

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LIST OF FIGURES

Figure page 1-1 The State of the Art of Artificial Organs .............................................................................. 17

1-2 A hybrid bionanodevice ......................................................................................................... 18

2-1 Microtubules and Dynamic Instability.................................................................................. 26

2-2 Kinesin motility on a microtubule......................................................................................... 27

2-3 In vitro motility assay ............................................................................................................ 28

2-4 Diagram of an epi-fluorescence microscope and spectra of a filter cube ........................... 29

2-5 Numerical aperture of an objective ....................................................................................... 30

2-6 Confocal Microscopy Schematic .......................................................................................... 31

3-1 Two approaches to protein photopatterning on thermoresponsive polymer layers using light-to-heat conversion in solution (A) and on the surface (B) ............................... 39

3-2 Photopatterning proteins using LHC in solution .................................................................. 40

3-3 LHC photopatterning of biomolecular motors on PNIPAM surfaces ................................ 41

4-1 Microtubule gliding velocities............................................................................................... 48

4-2 Microtubule gliding velocity as function of ATP concentration for a series of temperatures obtained by interpolating the data presented in Fig. 4-1 ............................... 49

4-3 Km (A) and maximal velocity vmax (B) as function of temperature..................................... 49

5-1 Light Activation and Control of Molecular Shuttles. .......................................................... 64

5-2 Measurement of shuttle velocity and radial distance from the center of illumination, and the experimental setup .................................................................................................... 64

5-3 Velocity profiles ..................................................................................................................... 65

5-4 Relationships between the maximum ATP concentration (A), the distance at which the ATP concentration drops to 10% of its maximum (B), and the control performance (C) and the characteristic distance according to eq. (1) and (2) .................... 66

6-1 Concept of a macroscale hybrid linear motor using nanoscale stators and forcers. .......... 73

6-2 Creating an isopolar microtubule array................................................................................. 74

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6-3 Analysis of isopolar microtubule arrays ............................................................................... 74

6-4 Force-generation of a microtubule array as a function of microtubule orientation. .......... 75

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

DESIGN, ASSEMBLY, AND OPERATION OF HYBRID BIONANODEVICES FOR

BIOMEDICAL APPLICATIONS

By

Robert Tucker

August 2009 Chair: Henry Hess Major: Materials Science & Engineering

Cells are the fundamental building blocks of life. Despite their simplicity, cells are

extremely versatile, performing a variety of functions including detection, signaling, and repair.

While current biomedical devices operate at the organ level, the next generation will operate at

the cellular level, combining the nanoscale machinery of cells with the mechanical robustness of

synthetic materials in the form of new hybrid devices.

This thesis presents advances in four topics concerning the development of nanomedical

devices: fabrication, stabilization, control, and operation. First, as feature sizes decrease from the

milli- and microscale towards the nanoscale, new fabrication methods must be developed. A new

rapid prototyping technique using confocal microscopy was used to produce freely-

programmable high-resolution protein patterns of functional motor proteins on thermo-

responsive polymer surfaces. Second, hybrid device operation should be temperature-

independent, but most biological components have strong responses to temperature fluctuations.

To counter operational fluctuations, the temperature-dependent enzymatic activity was

characterized for two types of molecular motors with the goal of developing a bionanosystem

which is stabilized against temperature fluctuations. Third, replacing electromechanical systems

consisting of pumps and batteries with proteins that directly convert chemical potential into

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mechanical energy increases the efficiency and decreases the size of the bionanodevice, but

requires new control methods. An enzymatic network was developed in which fuel was

photolytically released to activate molecular shuttles, excess fuel was sequestered using an

enzyme, and spatial and temporal control of the system was achieved. Finally, chemically

powered bionanodevices will require high-precision nano- and microscale actuators. A two-part

hybrid actuator was designed, which consists of a molecular motor-coated synthetic macroscale

forcer and a microtubule-based stator. Methods to create and characterize the stator were

developed, which can be used to optimize the force generation of the device.

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CHAPTER 1 INTRODUCTION

Within almost every cell exists complex nanoscale machinery, capable of repairing

damage,1, 2 communicating,3, 4 and creating entirely new cells with high fidelity.5, 6 The lifetime,

lifecycle, and morphology of a cell depend on the cell type and surrounding tissue.7-9 Some cells,

such as immune cells, operate autonomously, moving freely throughout the body to identify

foreign objects and pathogens.10-12 Other cells cooperate to better suit their tasks, working in

large groups to form tissues and organs.13

The human heart is one of the few organs to be successfully replaced with a permanently

implanted artificial device. Totally implantable artificial hearts were recently approved by the

United States Food and Drug Administration, but have limited lifetimes and duplicate only the

mechanical aspects of natural hearts.14-17 Although artificial hearts and ventricular assist devices

are considered successes, the devices are plagued by a variety of problems, including thrombosis,

encapsulation, and mechanical defects.18-20 Furthermore, while the heart has been traditionally

viewed as essentially a macroscale pump, there has been a recent shift towards a neuroendocrine

model, in which the heart responds to a variety of hormones, peptides, and chemical species

produced and released by the neuroendocrine system.13, 21-26 Although the heart as a whole

responds to these signals, it is the individual cardiomyocytes -heart muscle cells- comprising the

heart that detect these signals and alter their operating characteristics in response to the signals.

Without duplicating the cellular functionality of the cardiomyocytes, a true artificial replacement

for the heart cannot exist.

Similarly, the pancreas has been studied in great detail, but no permanently implantable

artificial device to duplicate its functions exists. The pancreas fulfills two roles within the body:

as part of the exocrine system it aids in digestion, and as part of the endocrine system it regulates

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blood glucose levels throughout the body. Specialized cell clusters within the pancreas called

“islets of Langerhans” produce insulin, a hormone that triggers the uptake of glucose into cells

throughout the body. Diabetes mellitus is a chronic autoimmune disease wherein the body can

no longer regulate blood glucose levels. This disease affects over 18 million people in the

United States alone.27 Despite extensive research, no cure for diabetes exists, and the current

treatments are crude. Diabetes is treated by monitoring blood glucose levels – either

continuously or more commonly several times throughout the day – and injecting insulin into

subcutaneous fatty tissue – either with syringes or a permanently attached external insulin

pump.28 While the macroscale glucose-regulating function of the pancreas has been mimicked

with multiple macroscale devices (Fig. 1-1 B), the cell-level and exocrine functions have not.

Implantable glucose sensors and insulin pumps will significantly reduce the impact of diabetes

on patient’s lives, but will not replace a pancreas. For this, a synthetic biomedical device capable

of replicating cell-level functions must be developed. Without duplicating the micro- and nano-

level characteristics of cells, fully functional replacement organs cannot be made.

Engineering at the Cellular Level

The challenges of engineering at the cellular level are significant. Cells use complex

processes to perform tasks such as cell motility,29, 30 inter- and intracellular communication,31, 32

and intracellular transport.33, 34 Cells move by the continuous polymerization and

depolymerization of actin filaments that are used to extend the periphery of the cell and

maneuver the rest of the cell behind it.35-37 Inter- and intracellular communication utilize small

chemicals, hormones, or proteins that are manufactured within the cell and released into the

surrounding area.38-40 Small molecules, if allowed to cross the cell membrane, can rapidly

diffuse through the cell to the target location.41, 42 In vivo, the mode of material transport

depends on the type and size of the material being transported.43 Larger molecules, proteins,

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and vesicles diffuse comparatively slowly, so cells use active transport to reduce the transport

time.44 Other cytoskeletal filaments, microtubules, provide structural support and function as

pathways for intracellular transport. Along these paths, molecular motors transport cargo,

working indivdually or in groups depending on the size of the cargo.45 This intracellular

transport system allows cells to detect and transport a variety of objects, independent of the size

or nature of the object.

The intracellular transport system can be integrated into hybrid bionanodevices that

combine the functionality of biological nanomachines with the ability to design and optimize the

properties of synthetic materials. Hybrid devices will require new methods of fabrication,

stabilization, control, and operation due to processing and operating constraints of both the

biological and synthetic components. The key feature of hybrid devices, however, is their

nanoscale functionality. Current state-of-the-art technologies can be used to fabricate

comparatively simple synthetic devices with nanoscale resolutions, i.e. junctions within

microprocessors, but complex patterns and functions are difficult to reproduce.46, 47 Hybrid

devices incorporate proteins which while difficult to control, already have nanoscale

functionality. Furthermore, control methods of synthetic devices have often focused on

controlling individual agents. By adapting cellular control strategies to hybrid devices, increased

functionality, higher efficiency, and better control may be possible. Finally, current technologies

have almost no ability to individually address nanoscale objects, a feat cells perform

continuously, i.e. proteins, vesicles, and filaments.

Firstly, proteins are extremely sensitive to environmental conditions including temperature,

ionic concentration, pH, and radiation.48-51 In contrast to the wide range of environmental

conditions used during fabrication of synthetic devices such as insulin pumps and electronics,

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fabrication of hybrid bionanodevices require new methods that are significantly closer to

physiological conditions. However, as device size decreases, new combinations of fabrication

techniques can be utilized which take advantage of the bottom-up self-assembly of biological

systems and the top-down engineering of synthetic systems. Many devices, such as biosensors,

use protein-protein interactions in their operation, and the ability to pattern proteins onto

artificial surfaces is of great interest.52-55 A rapid-prototyping method, termed light-to-heat-

conversion (LHC), was developed and used to pattern functional proteins onto thermo-

responsive polymer substrates using a confocal laser. Using LHC, multiple protein species can

were written directly onto a substrate with no loss in biological activity. In contrast to

photolithography that uses a fixed mask to pattern, the direct-write LHC procedure is freely-

programmable. Patterns with 2 µm resolutions were written and smaller resolutions are possible.

Secondly, biological systems respond strongly to temperature changes. Warm-blooded

animals use significant amounts of energy to maintain their body temperature, which allows

them to have comparatively simpler proteins that operate within a small temperature range.56

Small organisms and cold-blooded animals have enzymes and proteins which operate over a

wide temperature range, either due to different sets of proteins that operate at different

temperature ranges or more complex protein structures which compensate for temperature

changes.57, 58 Bionanosystems will be implemented in a variety of settings and temperatures, and

as such must have built-in operational stability against temperature fluctuations. The

microtubules in the bionanosystems presented are propelled over the surface by kinesin, a motor

protein present in many organisms whose specific activity depends on the host organism. The

activity of proteins is described using Michaelis-Menten kinetics, a common model for enzyme

kinetics which assumes that the reaction between the enzyme and substrate is thermodynamically

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driven and is at steady-state. Based on these assumptions, the enzyme can be characterized with

two values: 1) Vmax, the maximum velocity of substrate turnover and 2) KM, the substrate

concentration at which the turnover is one half of the maximum velocity and below which the

velocity is linearly dependent on substrate concentration. Kinesin from a thermophilic fungus

has been previously shown to have an increasing KM as temperature is increased, resulting in

constant activity across a wide temperature range.49 The activity of this kinesin was compared

with the activity of conventional kinesin derived from drosophila melanogaster in an attempt to

develop a temperature insensitive bionanodevices.

Thirdly, hybrid bionanosystems directly convert chemical energy into mechanical

movement, in contrast to many current biomedical devices that use electricity for activation and

control. Although non-electrical systems can be made smaller as no battery is required, new

control strategies must be developed. For example, electrically controlled devices can address

individual components, but hybrid devices must shift towards addressing swarms of components,

as found in cellular control strategies.59 A biomimetic approach to dynamically control hybrid

bionanosystems was developed wherein fuel was locally released to spatially and temporally

control system activation and excess fuel was subsequently sequestered.

Finally, macro- and microscale actuators have been produced primarily using a top-down

approach, although combinations of bottom-up self-assembly and top-down approaches have

been previously developed.60 In nature, actuators, such as muscles, are built using molecular

motors that work in a coordinated manner.45, 61 Although the force generated by individual

motors is on the order of piconewtons, large arrays can generate forces on the order of

kilonewtons. The force-generation by single or small groups of molecular motors has been

investigated in vitro in great detail, but biomimetic actuators based on large arrays of motor

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proteins have yet to be developed.62-66 A new method to create isopolar microtubule arrays was

developed and optimized to increase the force-generating capabilities as a function of

microtubule number, length, and orientation. Actuators driven by nanoscale motor proteins have

the potential for high efficiency and high precision that cannot be achieved without the

combination of top-down and bottom-up approaches.

Hybrid Bionanodevices

The intracellular transport system found within cells can be mimicked, augmented, and

integrated into artificial bionanodevices. Hybrid devices composed of synthetic substrates and

biological machinery have the potential to combine the versatility of immune cells and the

cooperativity of organs. For example, Figure 1-2 presents a concept for a three-chambered

hybrid biosensor which can be engineered to detect analytes, such as proteins and viruses. In the

first chamber the analytes enter the device and diffuse throughout the chamber. The particles

settle to the kinesin-coated surface where molecular shuttles composed of antibody coated

microtubules capture the analytes. The virus particles are transported to the second chamber

where they are bound to quantum dots via a second antibody, creating a double antibody

sandwich.67 The molecular shuttles loaded with virus particles, antibodies, and quantum dots are

transported to the third chamber which is illuminated with a light, causing the tagged viruses to

fluorescence, signaling analyte presence. The use of multiple binding steps (microtubules to

virus, viruses to quantum dots, etc) reduces the number of false positives of the biosensor. In the

case presented, only one analyte is being detected, but multiple types of antibodies and quantum

dots of with different emission wavelengths can be used to simultaneously detect a variety of

analytes.

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Figure 1-1: The State of the Art of Artificial Organs. (A) The heart has traditionally been viewed as a macroscale pump. However, recent research has shown heart muscle cells respond to and release a variety of signaling molecules. Artificial hearts are macroscale pumps that cannot replicate the intercellular communication of natural hearts. (B) A pancreas operates primarily at the cellular level, which current biomedical technology cannot duplicate. The current state-of-the-art artificial pancreas consists of several external devices, all of which operate at the macroscale. Adapted from 68-71.

A

B

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Figure 1-2: A hybrid bionanodevice. Analytes enter the device where they are collected and transported to a second chamber. The particles are tagged with antibodies specific to the analyte and fluorescently tagged. The tagged particles are transported to the collecting chamber where the signal is detected. Adapted from 72.

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CHAPTER 2 BACKGROUND AND EXPERIMENTAL TECHNIQUES

Knowledge and techniques from a variety of fields including material science, chemistry,

microscopy, biochemistry, physics, and biology were required to perform the experiments

discussed in the following chapters. The purpose of this chapter is to present background

material regarding the protein-based assays and describe in further detail the unique experimental

setups employed throughout the dissertation.

The Cytoskeleton

Cells contain a constantly changing proteinaceous infrastructure - the cytoskeleton - that

provides a means to control cell morphology, cell motility, and a variety of internal processes.

The cytoskeleton is composed of three classes of filaments: actin filaments, intermediate

filaments, and microtubules.73 Actin, with its complementary motor myosin, is the primary

component of muscles. While actin plays an active role in many cellular functions, its

cytoskeletal functions involve cell motility and counteracting tensile forces on the cell.

Intermediate filaments are found throughout the cell and are composed of tetramerized alpha-

helical rods, although the terminal domains of the rods, and thus their material characteristics,

depend on their intracellular location. For example, lamins are located in the nuclear membrane

to maintain the stability of the nucleus,74, 75 vimentin is found in the cytoplasm where it forms a

composite network with actin and microtubules,76 and desmin is found in muscle cells where it

forms part of the sarcomere.77 Microtubules, the largest of the three filament types, take part in a

variety of intracellular functions including intracellular positioning,78 intracellular transport,79

and counter-acting compressive forces.59

Microtubules are tubular structures that have an outer diameter of 24 nm and can be tens of

micrometers long.80 They have a three part structural hierarchy beginning with heterodimeric α−

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and β-tubulins that polymerize into protofilaments, which then combine to form microtubules.

Each tubulin monomer is approximately 4x4x4 nm3 with a molecular mass of 55 kDa (Figure 2-

1A).80 The protofilament and microtubule formation (Figure 2-1B, 2-1C) is powered by

hydrolysis of guanosine tri-phosphate (GTP), although the specific mechanism of polymerization

remains under investigation. Most microtubules consist of 13 protofilaments, although the

number in each microtubule can vary between 11 and 17.81-85 Microtubules were first discovered

in the 1950s and considered static structures,86, 87 but their continuous in vivo polymerization and

depolymerization – termed dynamic instability - was discovered fifteen years later by Inoue

(Figure 2-1C).88, 89 This dynamic instability allows the comparatively rigid microtubules to form

soft materials and rapidly respond to the ever-changing needs of the cell.90

Intracellular Transport

Cells use two internal transport methods, passive, which does not require an external

energy source and active, which does. Passive diffusion can be used to transport down

concentration gradients, over short distances, or when the diffusion constant is large, but is

inefficient over larger distances (i.e. multi-micron length-scales), within short time-frames, and

with small diffusion constants. Under these circumstances cells use active transport which,

while requiring energy, can be much faster than passive diffusive transport. During intracellular

active transport, microtubules and actin filaments are used as roadways that molecular motors

use to transport vesicles and other cargos throughout the cell.

Kinesin is a motor protein that converts chemical energy in the form of adenosine tri-

phosphate (ATP) into mechanical energy in the form of movement (Figure 2-2). Kinesin

consists of two regions: the heads which reversibly bind to the microtubule lattice and the tail

which binds to cargo. Each head contains an ATP-binding pocket, a 4 nm region which binds to

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the β-tubulin of the microtubule, a neck-region which chemomechanically couples the two

heads, and a coiled-coil tail region of variable length but is on the order of 70 nm. While

walking along a microtubule, the leading head is bound to ATP and the β-tubulin. The binding

of the leading head to the microtubule causes the rear adenosine di-phosphate (ADP) bound head

to swing forward 16 nm to the next β-tubulin. ATP-hydrolysis in the rear head releases ~20

kBT66, 91 of energy and forms water and ADP, causing ADP in the now-leading head to be

released. ATP then binds to the leading head, the rear head is ADP-bound, and the cycle begins

again. Each cycle takes roughly 1/100th of a second, resulting in a kinesin speed of 1000-1,700

nm/s under physiological conditions.92, 93

In-vitro Motility Assay

Within cells, microtubules form the “roadways” while kinesin acts at the cargo-carrying

shuttle.93, 94 In this work, the system was inverted, resulting in microtubules gliding over

surface-immobilized kinesin. The microtubule structure was stabilized with taxol, a chemical

that halts dynamic instability by stopping microtubule depolymerization.95, 96 The motility

assays take place in a “flowcell”, a microscope slide and slipcover separated by double-sided

tape, creating a chamber open at two ends through which solutions can be flowed (Figure 2-3A).

The motility assay is performed as follows: First, a flowcell is assembled and the inner

surfaces are passivated with a protein coating, usually casein or bovine serum albumin. Second,

motor proteins are flowed through which bind to the surface. Finally, a solution containing

microtubules, ATP, and antifade is added to the flowcell (Figure 2-3B). The microtubules

diffuse to surface, bind to the kinesin, and are imaged using epi-fluorescence microscopy (Figure

2-3C), which is discussed in the following section. In the in vitro motility assay, tubulin subunits

are tagged with a fluorescent rhodamine-dye, which when illuminated with green light (λ = 535

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nm), produces photons (λ ≈ 610 nm) and oxygen free-radicals. Without sequestration of the free

radicals, the sample will photobleach, a catch-all term used to describe the denaturation of the

dye molecules and the studied proteins, resulting in decreased fluorescence signal. To counteract

this phenomenon, a number of “antifades” have been developed to sequester the oxygen radicals

and extend the sample illumination time.97 The antifade used in the discussed work consists of

catalase, glucose oxidase, D-glucose, and dithiothreitol (DTT). The catalase converts water and

oxygen into hydrogen peroxide. The hydrogen peroxide is combined with D-glucose by glucose

oxidase to produce D-glucono-1,4-lactone. The DTT is a strong reducing agent that readily

donates electrons to the oxygen radical produced in fluorescence, thereby increasing the rate of

oxygen sequestration.

Wide-field Epi-fluorescence Microscopy

Although microscopy was invented a thousand years ago, a revolution in fluorescence

microscopy in biological research began when M. Chalfie and colleagues developed a technique

to express fluorescent proteins in living organisms in 1994.98-102 Fluorescence, a phenomenon

originally described by Sir George G. Stokes in 1852, is the term used to describe the absorption

of light of one wavelength by a compound and the subsequent emission of a lower energy, longer

wavelength of light.103

The experimental setup for wide-field fluorescence microscopy is shown in Figure 2-4A

and consists of four basic parts: 1) a light source, 2) a filter set, 3) an objective and 4) a sample.

White light is produced with a xenon or mercury arc-lamp and filtered to pass only visible

sections of the electromagnetic spectrum (350 nm < λ < 800 nm). Selection of the filter set

depends on the fluorescent dyes being used. Regardless of the specific fluorophores being used,

the set consists of three filters known as excitation, dichroic, and emission filters. The excitation

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filter is a band pass filter that passes a small subset of the full spectrum (λexcitation ± 25 nm)

produced by the arc-lamp, exciting the fluorophores within the sample (Figure 2-4B, blue line).

The excitation light reflects off the dichroic filter (also called a dichroic mirror) and is focused

by the objective into the sample. The dyes within the sample absorb the excitation photons and

release lower energy, longer wavelength photons (λemission). The emission light is collected by

the objective and passes through the long-pass dichroic filter (Figure 2-4B, green line) and band-

pass emission filter (Figure 2-4B, red line) to the camera or eye-piece. The emission filter passes

only light originating from the dye in the sample (λemission ± 35 nm) while blocking all other

wavelengths of light. Different filter sets can be used in conjunction with different fluorescent

dyes, which allows localization of different proteins within a single sample.

As shown in figure 2-4, the objective is used to focus excitation light and collect emitted

light. The best objective to use depends on the application and a variety of factors including the

numerical aperture (NA) of the objective, the magnification, and the resolution required. The

numerical aperture is a function of the refractive index of the sample media (n) and the half-

angle of the maximum cone of light that can enter or exit the lens of the objective (θ), as shown

in Figure 2-5. Air objectives (n = 1.0) are generally used for lower magnifications (40x and

below), and water (n = 1.33) or oil (n = 1.515 = nglass) immersion objectives are used when

higher magnifications (60-100x) are required. Oil immersion objectives have shorter working

distances (distance from the objective to the imaging plane) and are used to image objects near

the surface of the coverslip. In contrast, water objectives collect less light due to refractive index

mismatches, but have longer working distances and introduce fewer optical abberations, which is

useful when imaging in vivo processes. The resolution is limited by the illumination wavelength

and the numerical aperture, as shown in Equation 2-1 below, where r is the minimum resolution,

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λ is the wavelength of light used to illuminate the sample, and NA is the numerical aperture of

the lens.

λr (2-1)2 NA

=⋅

Confocal Microscopy

Confocal microscopy is a type of fluorescence microscopy that is identical to wide-field

epi-fluorescence microscopy with several key exceptions. A diagram of the confocal microscope

experimental setup used in Chapter 3 is shown in Figure 2-6. The light source for a confocal

microscope is a high-intensity laser, in contrast to the relatively low intensity arc-lamps used in

wide-field epi-fluorescence microscopy. The laser passes through a pinhole which eliminates

unfocused excitation light that would otherwise hit the sample. Using two mirrors (one each for

the x- and y-directions), the laser is rastered over the surface of the sample. The laser is focused

by the objective and strikes the sample. As in epi-fluorescecnce microscopy, the fluorescent

dyes in the illuminated region of the sample are excited and emit lower energy photons, some of

which are collected by the objective. After passing back through the objective, the light passes

through the dichroic mirror and another pinhole which eliminates unfocused emission light. This

light is passed through to either the CCD camera or the eye-piece.

The three main advantages of confocal microscopy are: (1) higher signal-to-noise ratios,

(2) higher temporal and spatial resolution, and (3) significantly less photobleaching of the

sample. Higher signal-to-noise ratios are achieved via a number of different methods. First, in

contrast to epi-fluorescence microscopy where large segments of the sample are illuminated,

only a small segment of the sample is illuminated at any one time in confocal microscopy. This

produces significantly less background light in the sample, thus a higher signal-to-noise ratio.

Secondly, the use of pinholes (or multiple pinholes depending on the type of confocal

25

microscopy being performed) means unfocused light never reaches the sample to unintentionally

illuminate areas and unfocused emission light never reaches the camera, reducing the noise in the

image. Finally, imaging the sample with a high-intensity laser allows for shorter illumination

periods that produce the same number of photons from the sample, resulting in higher signal to

noise ratios and spatial resolutions. Confocal microscopy also has an increase in temporal

resolution which is advantageous when imaging processes within live cells, e.g. intracellular

vesicle transport. The dwell time, the time each pixel is illuminated by the laser, can be

controlled down to the microsecond, in contrast to the millisecond timing available in epi-

fluorescence due to either the shutter speed or the camera response time. Photobleaching is

decreased in confocal microscopy because only a small region of the sample is being excited, in

contrast to epi-fluorescence microscopy where the entire field of view and all regions above and

below the focal plane are illuminated, producing oxygen free-radicals. Although the absolute

time the sample can be illuminated before photobleaching occurs remains the same, the relative

time the sample can be imaged is much longer because only small regions are illuminated at a

time.

Confocal microscopy does have two distinct disadvantages that can limit its use. As with

wide-field epi-fluorescence microscopy, the number of different fluorophores used to image a

sample is limited by the number of different excitation wavelengths that can be generated.

However, in contrast to the relatively inexpensive option of simply changing filter sets in wide-

field epi-fluorescence microscopy, confocal microscopy requires having multiple lasers (one for

each fluorophore), which can be prohibitively expensive. Secondly, wide-field epi-fluorescence

microscopy can be used to image the entire field of view, in contrast to confocal microscopy

26

where the imaging speed for the entire field of view is limited by the rastering speed. Despite

these limitations, confocal microscopy has proven invaluable to biological research.

Figure 2-1: Microtubules and Dynamic Instability. Microtubules in vivo continually undergo rapid growth and shrinkage. A) Microtubules are composed of heterodimeric proteins known as α- (light brown) and β-tubulin (dark brown), which bind GTP and GDP. B) The microtubules polymerize into linear protofilaments using GTP-hydrolysis. C) The GTP-bound protofilaments form sheets at the end of polymerizing microtubules, creating a tubular structure. D) During “catastrophe” (rapid depolymerization), the GTP-cap (red tubulin in C) is lost and the GDP-bound dimers form curved protofilaments, destroying the tubular structure. Adapted from 104.

A

B

C D

27

Figure 2-2: Kinesin motility on a microtubule. A) An ATP-bound kinesin head reversibly binds to the β-tubulin of the microtubule heterodimers. B) The binding of the leading ATP-bound head causes the rear ADP-bound head to swing forward 8 nm to the next β-tubulin. C) ATP-hydrolysis in the rear head causes ADP in the now-leading head to be released. D) ATP binds to the leading head, ADP is in the rear head, and the cycle begins again. Scale bar in (D) is 4 µm. Adapted from 105.

28

Figure 2-3: In vitro motility assay. (A) The motility assay takes place in a flowcell assembled from a microscope slide, two spacers made of double-sided tape, and a coverslip. This creates a chamber through which solutions can be flowed. (B) The glass surface is first passivated with casein, and then kinesin. The kinesin heads bind to microtubules in solution and walk towards the plus-end, resulting in net movement of the microtubule towards the minus-end. (C) The fluorescently labeled microtubules are imaged using epi-fluoresence microscopy and a CCD camera. Adapted from 106.

A B

C

+ end

Casein

Microtubule movementObjective

Slide

Spacers

Coverslip

Flow in

Flow out

29

Figure 2-4: Diagram of an epi-fluorescence microscope and spectra of a filter cube. (A) White light from an arc-lamp is passed through an excitation filter, which passes only light of certain wavelengths. The dichroic mirror reflects the light towards the specimen through the objective, causing the dyes to fluoresce. The emitted light passes back through the objective, passes through the dichroic mirror, through the emission filter, and finally to the eye-piece or camera. (B) The spectra of the filters used to image rhodamine-labeled microtubules.

ASample

Objective

Camera

Arc-lamp

Excitationfilter

Emissionfilter

Dichroicfilter

HQ535/50xQ565 LPHQ 610/75m

Tran

smis

sion

(%)

B

30

Figure 2-5: Numerical aperture of an objective. The numerical aperture is a function of the

refractive index of the media between the objective and the sample (n) and the half-angle of the maximum cone of light that can enter or exit the lens (θ). Oil immersion objectives are used to collect more light than air objectives, because less light is lost due to refraction at the air-glass interface.

NA = n*sinθ

Objective lensOil Airn=1.5 n=1.0

θ

SlideMediaCoverslip

Imaging Plane

31

Figure 2-6: Confocal Microscopy Schematic. Confocal microscopy is a type of fluorescence microscopy, but higher spatial and temporal resolution images can be produced relative to wide-field epi-fluoresence microscopy. The key differences are the use of a high-intensity laser to illuminate the sample and the use of two pinholes to block unfocused excitation and emission light.

Laser

Camera

Excitation Pinhole and Filter

Emission Pinhole and Filter

Dichroic

Objective

32

CHAPTER 3 FABRICATION: DIRECT-WRITE, RAPID-PROTOTYPING OF BIOFUNCTIONAL

PROTEINS ON THERMORESPONSIVE SURFACES

Introduction

Patterning functional proteins onto artificial substrates is of interest in the development of

nanotechnology, tissue engineering, biosensors, and cell biology.107-119 Towards this end, a

number of chemical patterning methods based on optical lithography,120-122 atomic force

microscopy (AFM),123, 124 printing techniques,125 chemical vapor deposition (CVD)126-128 have

been recently developed. While each of these methods provides particular advantages, a general

trade-off between spatial resolution, throughput, and maximum pattern size exists. For example,

AFM-based techniques can be used to place small numbers of functional proteins with

nanometer lateral resolution, but are limited to low writing speeds and small pattern sizes. On

the other hand, optical methods, such as the light-based activation of functional groups or ligands

on the surface,129 deserve particular interest because these techniques often offer sufficient

resolution combined with the potential for high through-put production. However, when based

on conventional lithography, expensive metal masks are needed and only predefined patterns can

be created. Moreover, the high-energy of the ultraviolet radiation (λ ~ 350 nm) often needed to

trigger the photoactivation of proteins or protein-binding molecules can be harmful for biological

species.130 Continuous illumination with light can also lead to the photogeneration of highly

reactive radicals - causing undesirable effects including protein conformational changes and loss

of biological function. Here, an alternative approach is reported based on the transient

application of visible light for the high-resolution patterning of planar substrates with functional

proteins. The new method allows the production of freely-programmable patterns of single and

multiple protein species.

33

This method is based on localized light-to-heat conversion (LHC) combined with a

thermoresponsive polymer surface capable of binding proteins and maintaining their

functionality. Previously it was shown that poly(N-isopropylacrylamide) (PNIPAM) could be

used to control the binding of proteins onto thermoresponsive surfaces.131,132 There, the

conformation of PNIPAM molecules in aqueous solution was switched in a spatially

unstructured manner between the collapsed state at T > 33°C (protein-binding conformation) and

the swollen state at T < 30°C (protein-repelling conformation). Here, the approach was extended

by using optical signals to generate heat in a highly-localized manner (Figure 3-1). To efficiently

convert light to heat, (i) solution-bound malachite green molecules133 (Figure 3-1A) or (ii)

substrate-bound black-ink layers (Figure 3-1B) were employed. In both cases, the surface-grafted

PNIPAM molecules collapsed locally and allowed proteins in solution to bind to the surface

exclusively in the illuminated areas. When the illumination was turned off, the heat quickly

dissipated and the PNIPAM molecules resumed their extended conformation and blocked the

surface against further protein binding. The swollen polymer chains also protected the patterned

proteins during consequent fluid exchanges, providing a means to sequentially pattern multiple

protein species on the same surface without the need for specific linker molecules or elaborate

surface preparation.

Preparation of PNIPAM Surfaces

PNIPAM layers were prepared via a two-step procedure similar to methods described

previously131 on glass coverslips (for solution-based LHC) or blackened mica plates (for

substrate-bound LHC). Glass cover slips (Corning) were first ultrasonically cleaned in

chloroform for 30 min, placed in hot piranha solution (3:1 (v/v) concentrated sulfuric acid and

30% hydrogen peroxide) for 1 h, and finally rinsed several times with high purity water. High-

grade mica substrates (Ted Pella, Inc.) were dip-coated with “edding T100” black ink (edding

34

International GmbH), dried at 120°C for 1 h and baked at 320°C for 2h in a vacuum oven. The

baking resulted in the formation of a stable black layer, which was insoluble in organic solvents.

Subsequently, a 1.5 nm thick layer of polyglycidyl methacrylate (PGMA, Mn = 84 000 g/mol,

synthesized by free radical polymerization) was deposited on top of the substrates by spincoating

a 0.02%(w/v) PGMA solution in chloroform and baked at 130 °C for 20 min in a vacuum oven.

After baking, a thick film (200 nm) of carboxy terminated poly(N-isopropylacrylamide)

(PNIPAM-COOH, Mn = 49 900 g/mol, PDI = 1.46, synthesized by anionic polymerization,

purchased from Polymer Source, Inc.) was spincoated on top of the PGMA layer from a 2%(w/v)

solution in chloroform and baked in a vacuum oven for 2 h at 180°C. Upon heating, the chemical

reaction between the terminating carboxyl groups of the PNIPAM and the epoxy groups of the

PGMA resulted in the formation of the grafted PNIPAM layer.131, 134-136 Non-grafted polymer

was removed using Soxhlet extraction in chloroform for 3 h. The thicknesses of the PNIPAM

layers (height = 6.5 - 7 nm) were determined by ellipsometry (SENTECH SE-402 scanning

microfocus ellipsometer, Sentech Instruments GmbH, Berlin) using a silicon wafer as a reference

sample (λ = 633 nm and 70° angle of incidence). The measured thicknesses corresponded to a

four-layer model Si/SiO2/PGMA/PNIPAM, where it was assumed that the polymer films

(PGMA and PNIPAM) had the same refractive index as the corresponding bulk polymers.

LHC-based Photopatterning of Proteins

To demonstrate the two LHC approaches, fluorescein-labeled casein (FITC-casein) was

patterned onto PNIPAM-coated surfaces using a confocal microscope. As the resolution limit of

photolithographical methods is directly related to the wavelength of illuminated light, UV light

was first used to achieve the highest resolution. Although UV has been shown to denature and

otherwise damage proteins, the initial experiments were conducted to test the efficacy of the

35

patterning technique, not the biofunctionality of the patterned proteins. In the first experiment, a

solution of FITC-casein (0.5 mg/ml) and malachite green (2.5 mM) in BRB80 buffer was

perfused into a 'flowcell' constructed from two coverslips (a 22 x 22 mm2 PNIPAM-prepared

coverslip and a 18 x 18 mm2 PEG-coated glass coverslip, Corning) and two pieces of double-

sided tape as spacers. The surface was then patterned by selecting areas on the PNIPAM surface

with imaging software (Zeiss-LSM AIM 3.2, Carl Zeiss GmbH, Germany) and illuminating them

with a confocal microscope (Zeiss LSM 510, Carl Zeiss GmbH, Germany) using a 99 mW UV

laser (λ = 351 & 364 nm, Enterprise UV, Coherent Inc, USA). The confocal microscope was set

to the maximum scanning speed (1.2 s to scan the full field of view, 7.2 µs/pixel dwell time) and

the pattern was illuminated 200 times, as this was found in previous experiments (data not

shown) to maximize protein binding to the surface while minimizing the pattern resolution.

After patterning the surface, non-adsorbed protein was removed by multiple perfusions with pure

BRB80 buffer, leaving the pattern of fluorescent protein adsorbed onto polymer layer. Using the

malachite green LHC patterning method, a variety of pattern shapes and sizes were easily

patterned (Fig. 3-1C). Using this method, a ~2 µm feature size was achieved, as seen by the line

widths shown in Figure 3-2. As the patterning method was based on confocal microscopy, any

pattern size or shape with a resolution limit of 2 µm can be written, not only lines or regular

shapes. However, malachite green is known to denature proteins upon heating,137 and can only

be used to pattern proteins whose biofunctionality is not required (e.g. casein is a globular

protein used primarily to block surfaces, whereas kinesin must retain its tertiary structure to

provide motility).

In the second experiment, the flowcell was constructed as above but with a blackened LHC

mica surface coated with immobilized PNIPAM, in place of the bottom coverslip (see Figure 3-

36

1D). The non-patterned proteins were removed by flushing the flowcell with pure BRB80. A

series of experiments was performed to clarify the mechanism of the fluorescent pattern

formation. First, it was found with the LHC layer absent, no pattern was created on the surface.

Second, when the thermoresponsive PNIPAM was substituted with non-thermoresponsive

polyethylene glycol (PEG), no patterning effect was seen. These two experiments demonstrate

that the combination of light-to-heat conversion and heat-induced change of the properties of

polymer layer are required for photopatterning.

Confirmation of the LHC Effect in Gliding Motility Assays

To test the efficacy of the LHC layer, gliding motility assays were performed directly on

the blackened LHC surfaces (without PGMA). In vivo, kinesin-1 ('kinesin')138, 139 is an

intracellular transport protein which carries cargo along microtubules, hollow, cylindrical protein

filaments 24 nm in diameter that span the interior of the cell. This system was inverted in the

gliding assay, and microtubules 5-10 µm in length gliding over substrate-bound kinesin. Because

the gliding speed of microtubules on kinesin surfaces was known to strongly depend on

temperature, it was possible to estimate the solution temperature when illuminated.140

Motility experiments were performed in a 5 mm wide flowcells constructed from a glass

cover slip (Corning, 18 x 18 mm2) and a mica plate (24 x 24 mm2) with LHC layer. A casein

solution (0.5 mg/mL casein in BRB80 (80 mM PIPES/KOH pH 6.9, 1 mM EGTA, 1 mM

MgCl2) was perfused into the flowcell and allowed to adsorb to the surfaces for 5 min. Next, 20

µL of a kinesin solution containing 2 µg/mL wild-type kinesin in BRB80 (full length drosophila

conventional kinesin expressed in E. coli and purified as described in ref 141) was perfused into

the flowcell and allowed to adsorb for 5 min. Finally, a motility solution containing rhodamine-

labeled taxol-stabilized microtubules,142 1 mM ATP, and an oxygen-scavenging system (20 mM

37

DTT, 0.02 mg/ml glucose oxidase, 0.008 mg/ml catalase, 20 mM D-glucose) in buffer was

perfused into the flowcell.

In this experiment, the bulk flowcell solution temperature was maintained at 7°C by means

of a Peltier element. When illuminating the fluorescently-labeled microtubules with short pulses

of green light (100 ms exposures with 1 s intervals, illumination with an HBO 100 (Osram) arc

lamp, excitation filter 480 nm, 40x air objective) measured microtubule gliding velocities were v

= 0.19 ± 0.04 µm/s (mean ± standard deviation, n=20 microtubules), which is within the

expected range for the given temperature.140 In contrast, when using long pulses (1 s exposures

with 100 ms intervals) the gliding speed increased to v = 0.7 ± 0.1 µm/s (n = 20 microtubules).

Control experiments on similar surfaces but without the layer of black ink did not show an

increased speed upon the long pulse excitation. Based on these gliding velocities, the

temperature of the kinesin was increased from 6°C to approximately 22°C. This experiment

provides reasonable evidence for conversion of light into heat on the LHC surface.

Testing the Functionality of Patterned Proteins

The functionality of the proteins patterned by substrate-bound LHC was tested using the

kinesin-microtubule system described above. Using a similar experimental preparation as for the

patterning of FITC-casein, a casein solution was perfused into the flowcell and incubated at 35°C

for 5 min. The flowcell was then cooled down to room temperature (25°C) and the kinesin

solution (10 µg/mL) was perfused in. A section of the blackened substrate surface was then

illuminated through a 100x objective (Zeiss, Plan-Neofluor, NA1.3, oil) using green light (λ =

480 nm, HBO 100 (Osram) arc lamp for 10 s). After the illumination was switched off,

microtubule-containing motility solution was perfused into the flowcell at room temperature and

the entire flowcell was heated to 35°C to collapse the PNIPAM and allow the microtubules to

38

land on the surface. As expected, microtubules landed on the patterned kinesin area and were

propelled over the surface. In contrast, no microtubules bound to the unpatterned casein surface

and those seen near the surface in solution were removed when washing the flowcell with

motility solution without microtubules. Consequently, the patterned circular area where

microtubules were continuously gliding at high temperature (35°C) became clearly visible

(Figure 3-3). Cooling the flowcell to room temperature led to release of microtubules due to

steric repulsion from the swollen polymer chains. This experiment clearly demonstrated (i) the

ability to pattern proteins with visible light and (ii) that the adsorbed proteins retain their

function and are not denatured.

In conclusion, a new approach was demonstrated to photopatterning functional proteins

based on their capture by thin films of thermoresponsive polymer locally heated by LHC

conversion (i) in solution and (ii) on a LHC surface. The LHC technique can use any wavelength

of light (UV to VIS) under physiological conditions. While the patterning of a single type of

protein at one time was demonstrated, the method can readily be used to pattern different protein

species by sequential cycles of light-induced adsorption. Although the LHC pattern resolution is

lower than AFM or CVD techniques, sub-micron resolutions should be attainable based on heat-

diffusion simulations. Furthermore, because the technique is maskless, freely-programmable

pattern sizes and shapes can be created. It is expected that this technique can find wide

application as a method of rapid prototyping for the fabrication of protein microarrays in bio-

and nanotechnological applications.

39

Figure 3-1: Two approaches to protein photopatterning on thermoresponsive polymer layers using light-to-heat conversion in solution (A) and on the surface (B). Localized illumination and subsequent conversion of light into heat causes the collapse of the thermoresponsive polymer, resulting in localized adsorption of proteins in the illuminated areas. When the illumination is removed, the photopatterned proteins remain entrapped in the polymer layer, and the swollen polymer chains prevent further protein binding. Epi-fluorescent images of FITC-casein surface pattern on the thermoresponsive polymer layer obtained by UV light (λ = 360 nm) laser beam irradiation using malachite green mediated light to heat conversion (C) and on blackened ink (D).

40

Figure 3-2: Photopatterning proteins using LHC in solution. A) FITC-casein patterned onto a PNIPAM surface using UV (λ = 351 & 364 nm) and green (λ = 633 nm) light. The lines are approximately 2 µm in width. (B) The graph is a linescan across the pattern (dashed vertical line (A)) comparing the pattern intensity with the background. For this experiment, the LHC material was malachite green in solution.

41

Figure 3-3: LHC photopatterning of biomolecular motors on PNIPAM surfaces. (A) Kinesin was photopatterned onto the immobilized PNIPAM layer which was grafted onto the LHC layer using green light (λ = 480 nm). After the patterning, the kinesin retained its biological functionality and provided continuous gliding of microtubules over the patterned surface at high temperature (35°C). Cooling down to room temperature (25°C) led to the release of microtubules due to steric repulsion of the swollen polymer chains. Fluorescent images show maximum projections derived from stacks of 10 epi-fluorescent images of microtubules released from the kinesin pattern (B) or gliding on the kinesin patterned surface (C).

42

CHAPTER 4 STABILIZATION: CHARACTERIZATION OF PROTEIN ACTIVITY FOR THE

DEVELOPMENT OF TEMPERATURE-INSENSITIVE HYBRID DEVICES

Introduction

A wide variety of nanodevices integrate biological components to provide unique

functions. One of the challenges arising from this hybrid approach is the stabilization of device

operation against temperature changes. The activity of biological nanomachines, such as

enzymes, is strongly temperature-dependent, often increasing 50 to 300% for every 10°C

increase in temperature at saturating substrate concentrations.143 Traditionally, temperature is

closely controlled in biotechnological processes and microfluidic devices,144-146 either to

maintain a stable activity or to switch between active and inactive states of the system. However,

in field-deployable devices it would be desirable to stabilize internal processes against

temperature fluctuations. Inspiration for stabilization strategies is provided by poikilotherm

organisms, who compensate for temperature changes on a wide range of timescales.56, 58

Two biological strategies providing instantaneous compensation are of particular interest.

On one hand, metabolic networks can display temperature compensation due to the existence of

feedback loops.147 The engineering equivalent is found in the canonical layout for temperature-

compensated electronic circuits.148 On the other hand, enzymes themselves can display near

constant activity for subsaturating substrate concentrations ([S]<Km) in a limited temperature

range. An example is lactate dehydrogenase from Alaskan king crab, whose Michaelis constant,

Km, increases with temperature and compensates for the increasing vmax value.57 From an

engineering point of view, this approach is preferable due to its simplicity and is similar to

employing a material with intrinsic temperature compensation such as INVAR steel.149

An interesting target for an exploration of this temperature compensation mechanism in the

context of hybrid devices is the ATPase kinesin. Kinesin and other motor proteins have evolved

43

in nature to provide actuation and transport within cells.150, 151 Recently, motor proteins and their

associated filaments (microtubules in the case of kinesin) have been employed for the transport

of nanoscale cargo in microfabricated structures,152-160 and it has been shown that these

“molecular shuttle” systems161, 162 can be applied to tasks such as force measurements,163 surface

imaging,164 single molecule manipulation,165 computing,166 and biosensing.67, 167, 168 Motor-

driven active transport is thus an attractive alternative to pressure-driven fluid flow and electro-

osmotic flow.169

Böhm et al.170 previously reported that the Km values for porcine kinesin-1 at temperatures

of 25°C and 35°C are 66±10 µM and 79±8 µM, respectively. Kawaguchi et al. determined that

the activation energy of bovine kinesin-1 is 50 kJ/mol, implying a Q10 at saturating ATP

concentrations equal to two.171 This suggests that the activity change per 10°C change in

temperature for small ATP concentrations is significantly smaller than two, implying that the

increasing Km partially compensates for the increasing vmax. Similarly, ATP consumption assays

for Thermomyces lanuginosis kinesin-3 determined an activation energy of 94±9 kJ/mol, and an

increase in the Km from 42±7 µM at 25°C to 1.6±0.3 mM at 50°C.49 At low ATP concentrations

the Q10 is 0.7 and thus the Thermomyces kinesin activity falls with increasing temperature

despite an increasing activity at saturating substrate conditions.

Here, for the first time, detailed measurements of the temperature dependence of kinesin

motor protein activity at subsaturating substrate concentrations from 19°C to 34°C for both,

Drosophila kinesin-1 and Thermomyces kinesin-3 is presented. These measurements were

undertaken in the expectation that - similar to other enzymes with applications in

biotechnology172 – the Km of kinesin would increase with increasing temperature with beneficial

implications for the design of kinesin motor powered hybrid devices.

44

Results

Velocity measurements for microtubules gliding on Drosophila kinesin-1 (Figure 4-1)

showed the expected Arrhenius-type increase with increasing temperature for a saturating ATP

concentration (1 mM) which replicates the data of Kawaguchi and Ishiwata171 for saturating ATP

concentrations (1 mM) obtained with bovine kinesin (see also 173).

A plot of velocity as function of ATP concentration for a series of temperatures (Figure 4-

2) was obtained by interpolating the data points in Fig. 1 for five temperatures for Drosophila

and four for Thermomyces. Fitting these curves with a Michaelis-Menten equation v([ATP],T) =

vmax(T) x [ATP] / (Km(T) + [ATP]) revealed the temperature dependence of the velocity at

saturating substrate concentrations vmax and the Michaelis constant Km (Figure 4-3). The 34°C

data point of the Thermomyces data was not included in the fit of vmax and Km since the reduced

vmax indicated partial deactivation.

The temperature-dependent vmax parameters were graphed in Arrhenius plots ln(vmax) =

vmax,0 x exp(-Ea/RT), and the activation energies were determined by linear error-weighted least-

square fits to be 53±5.1 kJ/mol (Q10=2.04) and 38±3.3 kJ/mol (Q10=1.67) for Drosophila

kinesin-1 and Thermomyces kinesin-3, respectively. Independent of temperature, the Km values

were found to be 70±10 µM and 220±90 µM for Drosophila kinesin-1 and Thermomyces

kinesin-3, respectively. Since the Michaelis constant was found to be independent of

temperature in the case of the two tested kinesins, temperature stabilization of enzyme activity

cannot be achieved at the expense of enzyme turnover by reducing the substrate concentration.

Discussion

It was not surprising that the measurements in the temperature interval from 19°C to 34°C

do not replicate the dramatic increase of the Km of Thermomyces kinesin-3 observed by Rivera et

al. in hydrolysis measurements at 55°C. The tested temperature range was well below the

45

thermal optimum for this organism. In addition, several changes in the properties of

Thermomyces kinesin-3 were observed at temperatures above 45°C.49 A second preparation of

Thermomyces kinesin-3 showed significantly altered motility (more interruptions of gliding

motion and more stuck microtubules) and a Km of 175 µM at 23°C, which indicated that

variations between preparations existed.

It was more challenging to reconcile the increase of Km for porcine kinesin-1 observed by

Böhm et al.170 with the observations of the Thermomyces and Drosophila kinesin. A possible

explanation for Böhm et al.’s observation is the depletion of ATP from the solution over time in

the absence of an ATP replenishing system, which caused an apparent increase of Km of

Drosophila kinesin as the cell was heated over the course of an hour in the initial experiments.

Only the utilization of an ATP regenerating system174 prevented this effect in the presented

experiments for both, Thermomyces and Drosophila kinesin.

It could be argued that even if kinesin’s Km would increase in proportion to vmax, the

compensation strategy would require a reduction of substrate concentration to a fraction of Km,

and consequently an undesirable loss in device performance. However, organisms routinely

operate enzymes at subsaturing substrate conditions,56 and even for hybrid devices it is far from

self-evident that the optimum activation corresponds to the maximum activation. For example, it

was found that a velocity of 200 nm/s represents an optimum for the loading of cargo onto

kinesin-driven molecular shuttles using biotin-streptavidin linkages.175 Similarly, INVAR steel

sacrifices mechanical properties for a low thermal expansion coefficient.

For the case of kinesin, it was concluded that stabilization against temperature changes will

require the design of a suitable enzymatic network, adding to the complexity of the device

similar to the recently demonstrated enhanced robustness of a DNA nanomotor.176

46

Materials and Methods

All chemicals were from Sigma-Aldrich (St. Louis, MO) unless otherwise specified.

Kinesin and microtubule preparation

A kinesin construct consisting of the wild-type, full-length Drosophila melanogaster

kinesin heavy chain and a C-terminal His-tag was expressed in Escherichia coli and purified

using a Ni-NTA column as in 141. Thermomyces lanuginosus kinesin was prepared as in 49.

Rhodamine-labeled tubulin (3.2 mg/mL, Cytoskeleton, Denver, CO) was polymerized at 37°C

for 30 min in BRB80 buffer (80 mM PIPES, 1 mM EGTA, 2 mM MgCl2, pH 6.9) with 1 mM

GTP (Roche Diagnostics, Indianapolis, IN), 4 mM MgCl2, and 5% DMSO, and subsequently

diluted 100-fold into BRB80 with 10 µM paclitaxel.

Variable temperature motility assay

A new flowcell was assembled for each ATP concentration tested, which consisted of a

square coverslip (22×22 mm, FishersFinest No. 1, Fisher Scientific), double-sided tape as spacer,

and a circular coverslip (15 mm diameter, No. 1, Warner Instruments Inc.). In the Drosophila

kinesin assays, the inner surfaces of the flowcell were incubated with a casein solution (0.5

mg/mL in BRB80) for 5 min, and then a Drosophila kinesin solution (10 nM in BRB80, 0.1

mg/mL casein, and the desired ATP concentration) for 5 min. A motility solution containing 3.2

µg/mL microtubules, casein (0.2 mg/mL), an antifade solution (20 mM D-glucose, 20 μg/mL

glucose oxidase, 8 μg/mL catalase, 10mM dithiothreitol (Bio-rad Laboratories, Hercules, CA)),

an ATP-regeneration system (2000 Units/L creatine phosphokinase, 2 mM creatine phosphate –

ref. 174) and ATP (10 µM, 25 µM, 50 µM, 100 µM, 200 µM or 1000 µM) was introduced, and

the edges of the flowcell were sealed with Apiezon® grease to minimize evaporation. In the

Thermomyces kinesin assays the procedure was identical except the flowcell was not incubated

47

with casein, the motility solution did not contain casein, and the ATP concentrations tested were

10 µM, 100 µM, 200 µM, 1000 µM, and 5000 µM. The flowcell was placed on the heat stage

(Model RC-20, Warner Instruments Inc.), the top coverslip coated with thermally-conductive

silver paste and covered with an aluminum plate, and the ensemble was fastened together with

screws. The temperature of the flowcell was set to the desired temperature (19°C, 23°C, 26°C,

31°C, or 34°C) and automatically regulated by a temperature controller (Warner Instruments

Inc., TC-324B), which used an electric current and a pair of 20 Ω resistors to maintain a constant

temperature. The microscope objective (Nikon 100X oil immersion) was heated by a resistive

wire (Nichrome 60, Pelican Wire Co.) powered with a DC regulated power supply (EXTECH®

instruments). The power supply was manually regulated to heat the objective to the same

temperature as the flowcell.

Measurement of temperature

The thermistor from the automatic temperature controller was inserted into the heat stage

near the aluminum plate on top of the flowcell. A second thermocouple was attached to the

objective and read using a multimeter. Thermocouple readings in the temperature range of

interest were calibrated against an alcohol thermometer.

Measurement of velocity

An Eclipse TE2000-U fluorescence microscope (Nikon, Melville, NY) with a 100X oil

objective (N.A. 1.4), an X-cite 120 lamp (EXFO, Ontario, Canada), a rhodamine filter cube

(#48002, Chroma Technologies, Rockingham, VT), and an iXon EMCCD camera (ANDOR,

South Windsor, CT) were used to image microtubules on the bottom surface of the flowcells. A

series of 10 images were taken for each ATP concentration and temperature, and the gliding

velocities of approximately ten microtubules were measured using image analysis software

(ImageJ v1.37c, National Institutes of Health, USA). The experiment was begun at room

48

temperature (T=19°C) and the flowcell was heated to each new temperature until motility

ceased. The error bars shown in the figures are the standard error of the mean of the velocity

measurements.

Figure 4-1: Microtubule gliding velocities. (A) Microtubule gliding velocity on Drosophila kinesin-1 as function of temperature for various ATP concentrations. Black stars are the data published by Kawaguchi and Ishiwata171 for 1 mM ATP. (B) Microtubule gliding velocity on Thermomyces kinesin-3 as function of temperature for various ATP concentrations.

20 24 28 320

500

1000

1500

2000

2500

1000 µM Kawaguchi 1000 µM 200 µM 100 µM 50 µM 25 µM 10 µM

Velo

city

(nm

/s)

Temperature (oC)20 24 28 32

5000 µM 1000 µM 200 µM 100 µM 10 µM

A B

49

Figure 4-2: Microtubule gliding velocity as function of ATP concentration for a series of temperatures obtained by interpolating the data presented in Fig. 4-1. Lines are fits to Michaelis-Menten functions with vmax and Km as parameters. (A) Drosophila kinesin-1, (B) Thermomyces kinesin-3.

Figure 4-3: Km (A) and maximal velocity vmax (B) as function of temperature. While Km does not change significantly as a function of temperature, the dependence of vmax on temperature is well-fitted by an Arrhenius equation. Drosophila kinesin-1 – full squares, blue; Thermomyces kinesin-3 – open triangles, red.

0 50 100 150 200 10000

250

500

750

1000

1250

1500

19 °C23 °C26 °C31 °C

Vel

ocity

(nm

/s)

[ATP] (µM)

34 °C

0 100 200 2500 50000

500

1000

1500

2000

34 °C 31 °C 26 °C 23 °C 19 °C

A B

18 20 22 24 26 28 30 32 34

500

1000

1500

2000

2500

vmax

= 8nm x 2.0x1011s-1 x e-6366K/(T(°C)+273K)

v max (n

m/s

)

Temperature (°C)

vmax

= 8nm x 1.1x109s-1 x e-4600K/(T(°C)+273K)

18 20 22 24 26 28 30 32 340

100

200

300

Km = 70 µM + (0.1 ±� 0.7) x (T(°C)-25°C)

Km (µ

M)

A

B

Km = 218 µM - (0.3 ±� 0.3) x (T(°C)-25°C)

50

CHAPTER 5 CONTROL: ADAPTING CELLULAR CONTROL STRATEGIES TO HYBRID

BIONANODEVICES

Introduction

A challenge for nanotechnology is the dynamic and specific control of nanomachines by

the user. Molecular shuttles, consisting of cargo-binding microtubules propelled by surface-

immobilized kinesin motor proteins, are an example of a nanoscale system which ideally can be

selectively activated at programmable locations and times. Here a biomimetic solution is

discussed, where activating molecules were delivered locally via photolysis of a caged

compound and subsequently sequestered in an enzymatic network. The controlled sequestration

of the activator not only created a rapid deactivation when the stimulus was removed, but also

sharpened the concentration profile of the rapidly diffusing activator. This improvement came at

the expense of a reduced efficiency in the utilization of the activator molecules, suggesting that

these nanosystems are most efficiently addressed as swarm and not as individuals. This work

represented a step towards transferring the cellular control strategies of molecular activation to

bionanotechnology.

Bionanotechnology is concerned with the utilization of biological components in

nanotechnology,177 which to a varying degree necessitates the use of biological engineering

approaches in a wider sense, for example in the use of self-assembly to create extended

structures. However, a striking accomplishment of nature is not only to create nanomachines and

weave them into larger structures, but also to control their spatial and temporal activation via

specific signals. This controlled activation is often achieved through the delivery of small

molecules, whose spatial and temporal distribution is shaped by the actions of multiple enzymes

releasing or sequestering the activating species. Examples include intracellular signaling via

calcium,178, 179 NAD(P)H,180 or cAMP.181

51

In contrast, the dynamic and controlled activation of specific nanomachines has been

addressed in a technological context primarily by making light-activation an integral part of the

design as in the light-driven synthetic motors based on rotaxanes or catenanes,182 or by designing

devices which can be individually activated with a highly specific fuel molecule.183 A new,

chemical approach is to exploit reaction-diffusion systems to locally change buffer conditions

and activate enzymes.184

Results

Here a biomimetic approach to dynamically control motor protein-driven

bionanodevices,185 in particular kinesin-driven molecular shuttles is presented.186 Molecular

shuttles consist of a surface patterned with stationary kinesin motors and cargo-binding

microtubules transported by the motors. Localized release and enzymatic sequestration of the

substrate ATP creates a spatially and temporally well-defined concentration profile, which in

turn lead to the controlled activation of a small number of molecular shuttles, as shown in Fig 5-

1. This approach significantly expanded the scope of previous work,162 which demonstrated that

repeated, step-wise activation of kinesin-driven molecular shuttles can be achieved by photolysis

of caged ATP in a solution of hexokinase without localization and on the timescale of minutes.

The necessity for this basic enzymatic network, arose from the rapid diffusion of ATP

(D=3x10-10 m2/s),187 which outpaced the movement of kinesin-driven molecular shuttles by a

factor of one hundred on the timescale of one second (or ten on the timescale of one motor step).

The presence of hexokinase in the solution limited the average distance an ATP molecule could

diffuse and lead to an increased spatial gradient of the kinesin activity.

The localization of the illumination with UV light and thus the photolysis of caged ATP to

a cylindrical region with a radius of either 15 µm or 25 µm lead to a dramatic improvement in

52

the temporal control over the shuttle activation. The shuttles immediately (<1s) responded to the

illumination by beginning to move. After the illumination ended, the shuttle velocity dropped by

1/e on a timescale of 10 s without sequestration by hexokinase to 1 s with hexokinase present.

Previously,162 uniform illumination of the cell led to deactivation on a timescale of hours to

minutes in the absence and presence of hexokinase, respectively. Of course, this improved

temporal control was a consequence of the rapid diffusion of the released ATP away from the

illumination zone and its dilution in the surrounding solution.

Repeated illumination with 250 ms pulses caused the shuttles to move a distance equal to

the distance traveled with a single pulse equal to the total duration of illumination, which implied

that the average translation per pulse could be tuned to less than 50 nm.

While the diffusion of the activating molecule, ATP, benefited the control of activation in

the temporal domain, the situation was reversed in the spatial domain.

The velocity of individual shuttles as function of their distance from the center of

illumination (Fig. 5-2A) to determine velocity profiles was measured. Since the velocity profiles

of pulsed and continuous illumination were very similar, the discussion focuses on the steady-

state between the release, diffusion and sequestration of ATP reached during extended

illumination. The velocity profiles corresponded closely to ATP concentration profiles, since

shuttle velocities were less than 20% of the maximum velocity at saturating ATP concentrations.

The velocity profiles (Fig. 5-3) showed an approximately exponential decay of the velocity

with increasing radius. As the hexokinase concentration in the solution was increased from 0 to

5,000 units/L, the accelerated sequestration of ATP lead to a reduction of shuttle velocity by a

factor of 30-50 and a sharpening of the concentration profile by a factor of 5-7. Changing the

53

radius of the illumination zone from 25 µm to 15 µm reduced the observed velocities to a third,

roughly in proportion to the area of the illumination zone (at constant light intensity).

To model the system, the diffusion equation was augmented with a generation term

describing the release of ATP (as a function of light intensity and size of illumination zone) and

a sequestration term describing the consumption of ATP by hexokinase, and solved numerically

for cylindrical symmetry (see methods section). A velocity profile was calculated from the

generated time-dependent ATP concentration profile using the Michaelis-Menten equation for

kinesin with Km = 25 µM and vmax = 1000 nm/s.188, 189 In the presence of hexokinase, continuous

illumination led to a steady state in less than 500 s. In the absence of hexokinase, the width of the

velocity profile increases with the third root of the illumination time and reaches 0.2 mm FWHM

after 200 s.

However, in order to obtain a good fit to the experimental data, the calculated velocity

profile (at the time of observation) had to be multiplied by a factor f, ranging from 1 for the

curve with the highest observed velocity (150 nm/s) to 0.05 for the curve with the lowest

observed velocity (1 nm/s). This was likely a result of slowing interactions between the multiple

kinesin motors transporting each microtubule at small ATP concentrations, an effect which was

previously experimentally observed188 and is also suggested by theoretical considerations.190

The excellent fit between the shape of the calculated and the measured velocity profile

proved that the model approximations (e.g. neglecting the hydrolysis of ATP by the kinesin

motors and the depletion of caged ATP in the illumination zone) were justified.

Additional insights into the process were obtained from analytically solving the steady-

state reaction diffusion equation with cylindrical symmetry outside the illumination zone. In the

present situation a characteristic diffusion distance r* was defined for the ATP molecules by r*2=

54

(KmD/A) where D=300 µm2/s was the diffusivity of ATP, Km=0.12 mM was the Michaelis

constant of hexokinase with respect to ATP, and A was the activity of the hexokinase solution.

The approximate solution191 of the reaction diffusion equation outside the illumination zone can

be written in terms of r* and the radius of the illumination zone ri as shown in Equation 5-1

where C0 is the ATP concentration at the boundary of the illuminated zone.

( )0 *exp (5-1)ii r rr

C Cr r

− = ⋅

Velocity profiles (Fig. 5-3) were calculated from Equation 5-1 using v=fvmaxC/Kmkin,

where f was the above mentioned factor accounting for slowing motor interactions, Kmkin was the

Michaelis constant for ATP consumption by kinesin, and small ATP concentrations were

assumed.

The experimental data were fit to the above expression for v outside the illumination zone

while the velocity within the zone was assumed to be constant and equal to v0=fvmaxC0/Kmkin.

The fit to the six experimental profiles utilized six values of C0 as free parameters while the value

of r* was obtained from the known values of Km, D and A (r* = 210 µm, 66 µm and 21 µm for

50, 500 and 5000 units/L of hexokinase, respectively), and the values of f were the same as in the

numerical simulations. The values of C0 obtained from the fits were 1.4 µM, 0.9 µM, 0.5 µM for

ri = 15 µm at 50, 500 and 5000 units/L hexokinase activity and 3.3 µM, 1.9 µM, 0.7 µM for ri =

25 µm at 50, 500 and 5000 units/L hexokinase activity, respectively.

By equating the rate of ATP consumption integrated over all radii with the rate G of ATP

generation from caged ATP, the ATP concentration C0 in the illumination zone and at the

boundary could be determined. At low light intensities, the ATP generation was not limited by

the diffusive influx of caged ATP into the illumination region, but was given by G =

55

CcATP×k×Ihν, where CcATP was the constant concentration of caged ATP, k was the ATP uncaging

rate constant and Ihν was the UV light intensity.192 This resulted in the concentration formula

shown in Equation 5-2.

0 2

*

(5-2)1 1

cATP hv

i

C k IC

Drr

⋅ ⋅=

+

Based on the known values for CcATP, D, r*, ri, k and the fit values for C0, this implied that the

average illuminating UV intensity is 2 mW/cm2 for both pinhole sizes.

The analytical solution illustrated the opposing trends in achieving localized control: A

reduction in the characteristic ATP diffusion distance r* (increase in hexokinase activity) led to

reduced ATP levels (Fig. 5-4A) but at the same time narrowed the ATP plume as measured by

the radius at which the shuttle velocity was reduced to 10% of the maximum (r10%, Fig. 5-4B).

Interestingly, if the absolute concentration gradient C0/r10% was used as the performance metric

of the control process (Fig. 5-4C), it became apparent that an optimum for the characteristic

diffusion distance existed.

Since r10%≈ri+r* for r*<10ri (Fig. 5-4B), by finding the maximum of C0/(ri+r*) the optimal

value of the characteristic diffusion distance r* could be determined to be approximately equal to

twice the illumination radius ri (Fig. 5-4C).

Discussion

This suggested a strategy to achieve a desired level of control in the case of cylindrical

symmetry: (1) Define the size of the activation zone given by r10%, (2) restrict illumination to a

third of the activation zone ri=r10%/3, (3) adjust the activity of the sequestration enzyme so that

the characteristic diffusion distance equals twice the radius of the illumination zone (r*=2ri), (4)

56

tune the concentration of caged ATP and the intensity of illumination to achieve the desired

speed within the illumination zone.

Unfortunately, even in the optimal case a decrease in the size of the activation zone led to a

linear decrease of the generated gradient and a quadratic decrease in fuel efficiency, which was

defined here as the ratio of the number ATP molecules used by motors to the total number of

ATP molecules released. The decreasing gradient could be compensated by increasing the

intensity of the light source, but the amount of ATP which could be released was ultimately

limited by the influx of caged ATP into the illumination zone. However, the light intensity at this

limit was expected to lead to unacceptable levels of photodamage.

The above described experiments, simulations, and calculations demonstrated a significant

improvement in the control of molecular shuttles, however they also point out a trade-off: While

the sequestration of small molecules which govern the activation of nanosystems improved the

spatial and temporal control, even a restriction of the activation zone to tens of micrometers

reduced the activation level drastically. At the same time the vast majority of the released

control molecules did not even interact with the nanomachines. These general considerations

apply in a variety of experimental situations. For example, small molecules may be delivered by

injection into a fluid stream which rapidly removes them from a stationary target193. Metal ions

may control the activation of a restriction enzyme184 or F1-ATPase194 and be sequestered by a

chelator, and DNA motors195, 196 are controlled by DNA oligomers which may in turn be

removed by other oligomers. In the context of molecular shuttles, the release of cargo can be

achieved by delivering a competitor for the cargo linkage.197, 198

Alternatives to control via activating molecules exist but are not superior. Localized

heating in a cold environment146, 199 has been demonstrated, however for kinesin motors a ten-

57

fold difference in activation (chosen here as benchmark to define the activation zone) is difficult

to achieve due to the limited range of protein stability.171 Similarly, optical molecular switches

can overcome diffusion limitations, but currently only a roughly five-fold difference in activity

between the on- and off-state of the controlled nanosystem has been demonstrated.200, 201

Electronic switching of enzyme activity is promising, if the currently achieved on/off ratio of 3/2

can be improved.202 Control by uncaging and sequestration of control molecules, in contrast,

could potentially be increased up to the diffusion limit by employing a more brilliant light source

(e.g. a UV laser) and minimizing photodamage. A complementary approach to control is

“steering”, where fluid flow, electric, and magnetic fields are applied to orient actin filaments or

microtubules.203-208 In this case motor activity is not targeted. Similarly, switching the properties

of a surface to promote/inhibit the ability of adhered motors to bind microtubules controls

microtubule density rather than motor activation.209

These findings reiterate that due to the rapid diffusion of information in integrated

molecular factories, as exemplified by cells, spatial organization of the workflow is not efficient

– in contrast to the macroscale. Instead, information has to be stored in and manipulated between

a multitude of different chemical species. While it was shown that activation can be successfully

constrained by local release and rapid sequestration, it may be more natural to employ molecular

shuttles as parts of a large and dispersed swarm rather than individually. To quote Hofstadter’s

anteater, “… you must not take an ant for the colony”.210 The engineering task is then to tailor

the swarm behavior, e.g. by limiting dispersion169 or by self-generating ATP157 rather than the

individual nanotransporter.

58

The insights generated from this investigation of control via diffusing molecules may also

inform work at the interface of BioNEMS and systems biology,211 in molecular computing,212

and in the emerging field of bio-nano-logistics.213


Kinesin and Microtubules

Full length, wild-type kinesin from Drosophila melanogaster expressed with a C-terminal

Histidine-tag in E. coli was purified using a Ni-NTA column as described in ref 141.

Microtubules were polymerized as follows: Polymerization buffer (80 mM Pipes, 1 mM EGTA,

5 mM MgCl2, 1 mM guanosine 5’-triphosphate, and 5% dimethylsulfoxide) was added to a 20

µg aliquot of rhodamine-labeled tubulin (TL331M, Cytoskeleton, Denver, CO) resulting in a

final tubulin concentration of 32 µM. The solution was kept at 37°C for 30 minutes and then

diluted 100-fold in BRB80 (80 mM PIPES, 1 mM MgCl, 1 mM EGTA, pH 6.9) containing 10

µM taxol for stabilization and kept at room temperature (20°C).

Caged-ATP and Hexokinase

Photocleavable 1-(4,5-Dimethoxy-2-nitrophenyl)ethyl caged ATP (Molecular Probes,

Eugene, Oregon) was dissolved in nanopure H2O to a final concentration of 10 mM.

Hexokinase (H-5625, Sigma-Aldrich, St. Louis, MO) was dissolved in BRB80 to create

stock solutions of 1000-400,000 Units/L. These stock solutions were then diluted in motility

solutions resulting to final concentrations of 25 – 5000 Units/L. The KM for hexokinase with

respect to ATP is 120 µM.

Motility Assays and Microscopy

The motility assays were performed in 100 µm high, 1.5 cm wide flowcells. Casein (0.5

mg/mL, Sigma-Aldrich, St. Louis, MO) dissolved in BRB80 was adsorbed for 5 min to reduce

59

denaturation of kinesin. Kinesin (10 nM kinesin, 1 mM MgATP, 0.5 mg/ml casein, BRB80) was

then adsorbed for 5 min. Finally, the motility solution containing the caged ATP was flowed

into the flowcell (32 nM microtubules, 500 µM caged-ATP, and an oxygen-scavenging system

consisting of 20 mM D-glucose, 0.02 mg/ml glucose oxidase, 8 µg/ml catalase, 1% dithiotreitol

in BRB80). D-glucose is simultaneously the substrate for glucose oxidase and hexokinase.

Hexokinase hydrolyzes ATP while phosphorylating D-Glucose, producing ADP and glucose-6-

phosphate. Since the KM of hexokinase for glucose was 120 µM, the hexokinase activity was

limited only by the ATP concentration.

The flowcells were imaged using an epifluorescence microscope (Eclipse TE2000U,

Nikon) with a 100 W Hg lamp, a 40x oil objective (NA 1.4), a cooled CCD camera (Andor iXon,

Andor Technology, Windsor, CT) and a rhodamine filter set (Filter Set 48002, Chroma

Technology Corp, Rockingham, VT). The ultraviolet illumination in the wavelength range from

325 nm to 375 nm was provided by a xenon arc lamp (Lambda-LS, Sutter Instrument, Novato,

CA) with a liquid-light guide and passed through circular pinhole with a radius of 150 or 250 µm

(Edmund Optics Inc.) and a UV filter (Chroma D350/50x, Chroma Technology Corp,

Rockingham, VT). A 10x demagnified image of the pinhole was projected by the condenser

(High N.A. Condenser, Nikon Instruments) to the object plane of the microscope. The aperture

stop of the condenser was almost completely closed to create a nearly cylindrical light path (Fig.

5-1) with a conical angle of 6°.

The light passing through the flowcell was measured using a hand-held power meter

(Model #3803, New Focus, San Jose, CA) calibrated with a UV sensing power meter (Mannix

UV-340 Light Meter, Ambient Weather, Tempe, AZ). Within the flowcell, this light power of

0.8 µW and 1.3 µW was focused into a near cylindrical cone with a radius of 15 and 25 µm

60

resulting in an intensity of 110 and 65 mW/cm2, respectively. These intensities were fifty-fold

higher than the intensity implied from the analytical and numerical fits to the data. This

discrepancy was not resolved.

All assays were performed at 20°C.

Velocity and radius measurements

The image acquisition settings were chosen according to the velocity of the shuttles. The

time between image acquisitions for both pinholes was 10, 10, 40, and 80 s for hexokinase

activities of 0, 50, 500, and 5000 Units/L, respectively. The exposure time for all images was

500 ms.

For steady-state experiments, the UV source was “on” continuously throughout the

experiment and velocity measurements were begun after a minimum of 100 s of UV illumination

and extended up to 1000 s. The radial distance was measured from the center of the pinhole to

the leading edge of the microtubule in the initial image (blue arrow, Fig. 5-2). The velocity was

given by the ratio of the distance between the leading edge of the microtubule in the two images

(white arrow) divided by the time between images (200 s in Fig. 5-2). Images were skipped to

increase the accuracy at low velocities.

For the pulsing experiments, the UV source was on for 250 ms and off for 1,750 ms, for a

total cycle time of 2 s. 1000 pulsing cycles were performed for each hexokinase concentration

and pinhole, and the velocity and radius measurements were then derived from these images.

The shuttle velocity was calculated by dividing the distance traveled by the on time.

Numerical model

Equation 5-3 is a second order partial differential equation in cylindrical symmetry and

was solved for the ATP concentration within the flowcell using the ODEsolve routine in

MathCAD 13.1 (Parametric Technology Corp., Needham, MA).

61

2 2

2 2m

-[ATP] [ATP] D [ATP] [ATP] D G(r,t) A (5-3)

t r r r K [ATP]∂ ∂ ∂

= + ⋅ + ⋅∂ ∂ ∂ +

Here, the ATP concentration depended on the radial diffusion of the ATP (first two terms),

the ATP generated within the pinhole (third term), and sequestration by hexokinase (fourth

term). The ATP concentration was assumed to be zero initially and at a 1 mm distance from the

center of the flowcell for all times. The measured light intensity was used to calculate a first ATP

uncaging rate G(r,t) within the cylindrical illumination zone using previously determined

photolysis rates. The generation function, G(r,t) was set to a constant value within the

illuminated region (15 or 25 µm ,“on”) and set to zero outside of the illumination radius (“off”).

The resulting velocity was tuned to fit the experimental data shown in Figure 5-3 by varying the

f parameter shown in Equation 5-4. The Km for hexokinase was set to 120 µM.

The resulting ATP concentration profile was then used to calculate the molecular shuttle

velocity assuming Michaelis-Menten kinetics of kinesin with a vmax of 1000 nm/s, a Kmkin of 25

µM, inhibition constants (Ki) for ADP and caged-ATP of 35 µM and 200 µM and is shown in

Equation 5-4.

maxkin kin

kin m mm caged-ATP ADP

i i

V [ATP]V f (5-4)K K[ATP] K [caged-ATP] [ADP]

K K

=+ + ⋅ +

The factor f accounts for the decreasing shuttle velocity at small ATP concentrations and

was chosen as 0.75, 0.7, 0.6, 0.4, 0.15, 0.05 for shuttle velocities at the center of illumination of

65-100, 60, 40, 20, 6, 1 nm/s, respectively. The time to reach steady-state for constant UV

illumination varied depending on the pinhole size and hexokinase concentration, but steady state

was always reached within 500 s.

62

Analytical Model

The analytical solution for the steady state ATP concentration was obtained under the

assumptions that the ATP concentration inside the illumination zone was constant, and that the

consumption of ATP by kinesin compared to hexokinase was negligible. The steady state

diffusion and sequestration of ATP outside the illumination zone (assuming cylindrical

symmetry) is shown in Equation 5-5. In Equation 5-5, D is the diffusion constant of ATP, A is

the activity of hexokinase and Km is the hexokinase Michaelis constant for ATP

2

2

1 0 (5-5)m

C C AD Cr r Kr

∂ ∂+ − = ∂∂

Equation 5-5 can be rewritten as shown in Equation 5-6, where r*2 = Km/DA.

2 22

2 2 0 (5-6)*

C C rr r Crr r

∂ ∂+ − =

∂∂

Equation 5-6 is the modified Bessel equation, for which one of the solutions is shown in

Equation 5-7: 191

0 (5-7)*

rC aKr

=

In Equation 5-7, K0 is the modified Bessel function of the second order and a is an integration

constant. At the boundary of the illumination zone ri, the concentration C(ri) is equal to

concentration C0 inside the illumination zone. Applying these boundary conditions, the

concentration as a function of radius is shown in Equation 5-8.

00

0

(5-8)*

*i

C rC Kr rKr

=

K0(x) can be approximated as (1/x)*exp(-x), which reduces Equation 5-8 to the form shown in

Equation 5-9, which is identical to Equation 5-1.

63

0( )

exp (5-9)*

i ir r rC C

r r− =

64

Figure 5-1: Light Activation and Control of Molecular Shuttles. A nearly cylindrical cone of UV light is produced within a flowcell to locally photolyze caged ATP. As the ATP diffuses outwards, it is hydrolyzed by kinesin, resulting in localized microtubule movement. Hexokinase is added to the solution to increase the gradient of the ATP concentration profile. As the hexokinase concentration is increased, the area of activation and the maximum shuttle speed decrease.

Figure 5-2: Measurement of shuttle velocity and radial distance from the center of illumination, and the experimental setup. A) Two images (pseudo-colored in green and red and separated by 200 s in time) were overlaid, showing the illumination zone and the movement of microtubules with radius-dependent velocity, due to the ATP sequestration by hexokinase in solution (50 units/L). B) The experimental setup.

High [ATP]

Low [ATP]

V

r

microtubule

kinesin

ATP gradient

UV

illumination

65

Figure 5-3: Velocity profiles. Velocities of individual shuttles were measured as a function of the distance from the center of illumination for illumination zones with a radius of 25 µm and 15 µm at different hexokinase concentrations (0, 50, 500, 5000 units/L). Dashed lines are the numerical solutions and solid lines are the analytical solutions as described in the text.

0

25

50

0 50 100 150 20001

0 Units/L 50 Units/L500 Units/L

Spee

d (n

m/s

)

Radius (µm)

05

5000 Units/L

15 µm Radius of Illumination

5000 Units/L

25 µm Radius of Illumination

0

50

100

150

0 Units/L 50 Units/L500 Units/L

66

Figure 5-4: Relationships between the maximum ATP concentration (A), the distance at which the ATP concentration drops to 10% of its maximum (B), and the control performance (C) and the characteristic distance according to eq. (1) and (2). The broken vertical lines indicate the hexokinase activities used in the experiments with illumination zones of radius 15 µm and 25 µm.

67

CHAPTER 6 ACTUATION: ASSEMBLY OF AN ISOPOLAR MICROTUBULE ARRAY TO PRODUCE

MACROSCALE FORCES USING NANOSCALE COMPONENTS

Introduction

Bionanotechnology is a field that covers many topics, one of which is the integration of

biological components into engineered synthetic environments. Nature has proven that

nanoscale machines can be produced, as evidenced by cells and sub-cellular components. The

proteins found within cells have been optimized over billions of years of evolution, but utilizing

them outside of their natural environment has proven a challenging task. Hybrid devices have

been developed with proteins functioning as nanoscale machines placed into engineered

environments, resulting in devices and systems which have the nanoscale functionality found

within cells and the optimizability of engineered systems.214-217

Motor proteins such as kinesin and myosin have been studied in great detail, both in vivo

and in vitro.62, 65, 218-223 While there has been much investigation into the nanoscale force

generation by myosin, to date there have been no successful attempts to produce large-scale

forces using arrays of myosin, as seen in muscle.29, 30 Kinesin and myosin have been used to

transport micro- and nanospheres, but macroscale movement has yet to be achieved.175, 224, 225

Large-scale force generation is achieved in vivo via a complex hierarchical structure wherein

myosin motors and actin filaments form sarcomeres, sarcomeres combine to form myofibrils,

myofibrils then form muscle fibers, which finally assemble into muscles. The hierarchical

structure of muscle has many benefits,226 but is difficult to reconstitute given the current state-of-

the-art of fabrication methods. However, even at its lowest level of complexity, the sarcomere

can be viewed as a simple linear motor that has been optimized based on the material limitations

of its components and the requirements of the motor. These optimization parameters include the

length of the myosin motor and actin stator, length of the motor unit (sarcomere), the density of

68

motors, and the orientational order of the components. From these design characteristics, it was

learned what parameters are important in the design of a hybrid linear motor. An engineered

microtubule array is presented that can be optimized to maximize the force generation of a

hybrid linear motor. The motor will be powered by tri-phosphate (ATP) hydrolysis and will

consist of two parts: 1) A substrate-bound isopolar microtubule array as a stator and 2) a

macroscale object coated with kinesin, with the kinesin acting as the forcer. The combination of

these two components can produce macroscale forces and displacements in a controlled manner

through the use of a caged fuel source to control actuator activity. A concept of the linear motor

is shown in Figure 6-1. Multiple layers of the stators and forcers can be combined to produce

larger forces and displacements than with single layers. Furthermore, the combination of

different species of motors (i.e. dynein and kinesin) within the layers or anti-parallel orientations

of the arrays can be used for fine control the movement of the linear motors, or even produce bi-

directional movement (Figure 6-1B).

Results

Microtubules were bound to a kinesin-coated surface, aligned with fluid flow to produce

an isopolar array, and subsequently fixed to the surface using glutaraldehyde. The microtubules

land with random orientations, and the flexibility of the kinesin motors allow microtubules to

move in any direction over the kinesin surface.227 Although the overall microtubule structure is

stiff (Lp ≈ 5 mm),228 the leading edge of the microtubule is an order of magnitude more flexible

(Lp ≈ 100 µm)229, 230 and the movement of the tip can be biased using low shear forces. Under

flow, the lowest energy state of the microtubule is to move in the direction of and align parallel

to the direction of fluid flow. Under these conditions, the tip of the microtubule will still

fluctuate, but the movement of the tip motion will be biased to remain parallel to the direction of

69

flow, resulting in an isopolar microtubule array. The limitations of this alignment method are: 1)

the flow rate must be high enough to reorient the microtubule tips and 2) the flow must be

applied long enough for the microtubules to complete a turn to align with the flow direction

(where the alignment time is proportional to the length of the microtubule divided by the velocity

of the microtubule). Glutaraldehyde cross-links lysine residues between tubulin subunits and

stabilizes the microtubule structure, but at high concentrations, the ability of kinesin to bind to

the microtubule lattice is impacted.231 The glutaraldehyde concentration was chosen to securely

fix the microtubules to the surface while allowing new kinesin to walk along the microtubules, as

evidenced by successful bead-type assays (data not shown) and previous research.232, 233 The

beads walked in the same direction along different microtubules, showing the isopolar alignment

of the microtubules. Figure 6-2 shows the microtubules with random orientation upon landing

(Figure 6-2A), after alignment (Figure 6-2B), and after fixation with glutaraldehyde (Figure 6-

2C).

This project focused on the fabrication and characterization of the microtubule array.

Microtubule arrays were developed previously by several research groups, but no means of

quantitatively comparing the arrays exists.204, 234-238 The focus of this work was on the alignment

of the microtubules, as measured by the orientational efficiency as defined in Equation 6-1. In

essence, the efficiency is a measure of the average orientation of the microtubules within the

microtubule array compared to an array with perfect alignment. In Equation 6-1, ηorientation is the

orientational efficiency, N is the number of microtubules in the array, and α is the angle of the

microtubule relative to the direction of fluid flow (α=0°) in Figure 6-2.

orientation1

1 cos cos (6-1)N

Nη α α= =∑

70

Discussion

An example of the orientational distribution before and after alignment is shown in Figure

6-3A. Prior to alignment (red bars), the microtubule orientation was essentially random. After

alignment (blue bars), a majority of the microtubules were aligned within 10° of the direction of

fluid flow, and the orientational distribution had an approximately exponential decay. The

average orientational efficiency of four arrays before (68 ± 30%, mean ± stand. dev) and after

(94 ± 13%) alignment is shown in Figure 6-3B. As expected, both the efficiency and the

variation of the efficiency decreased with the alignment process. The flow rate used to align the

microtubules was varied between 2-6 µl/s, although variation in flowcell widths introduced

difficulties in directly linking flow rates and orientational efficiency. The shear rate was 600/s

and assumed a parabolic flow profile. The relationship between force generation of the arrays

and microtubule orientation within the array is shown in Figure 6-4. Within the non-linear

region of the plot (0.8 < η < 1.0), only modest gains in the force generation capabilities of the

array are made when increasing the orientational efficiency. Therefore, the optimum

orientational efficiency of the array should be above 0.8, but only minimal improvements in

force-generating capabilities will be gained above this level.

These insights can be used to create a model to determine the force-generating capacity

of a particular microtubule array, which in turn can be used to optimize array characteristics for

specific application. A new metric was proposed, termed “Linear Orientational Array Density”,

to characterize filament-based arrays. As shown in Equation 6-2, the linear orientational array

density (λ) is a function of the orientational efficiency (η), the microtubule surface density (σ),

and the average length of the microtubules comprising the array (<L>).

= L (6-2)λ η σ⋅ ⋅

71

Based on this model, the force generating capability of the array can be optimized prior to

completion of a fully functioning device. Future work will also focus on creating conformal

contact between the forcer and stator to maximize force generation of the motor.


Kinesin and Microtubules

Full length, wild-type kinesin from Drosophila melanogaster expressed with a C-terminal

Histidine-tag in E. coli was purified using a Ni-NTA column as described in ref 141.

Microtubules were polymerized as follows: Polymerization buffer (80 mM Pipes, 1 mM EGTA,

5 mM MgCl2, 1 mM guanosine 5’-triphosphate, and 5% dimethylsulfoxide) was added to a 20

µg aliquot of rhodamine-labeled tubulin (TL331M, Cytoskeleton, Denver, CO) resulting in a

final tubulin concentration of 32 µM. The solution was kept at 37°C for 30 minutes and then

diluted 100-fold in BRB80 (80 mM PIPES, 1 mM MgCl, 1 mM EGTA, pH 6.9) containing 10

µM taxol for stabilization and kept at room temperature (20°C).

Microtubule Array Preparation

Microtubules were aligned with an isopolar orientation using fluid-flow and subsequently

fixed to the kinesin surface. A flow cell was made by placing two microscope slip covers (35 x

75 mm2 and 22 x 22 mm2, thickness 0, FischersFinest, Fisher Scientific) with double-sided tape

as spacers, creating a chamber through which solutions were flowed. The inner surfaces of the

flow cell were passivated with 0.5 mg/ml casein in BRB80 buffer (5 min adsorption time) and a

~10 nM kinesin solution containing 0.2 mg/ml casein, and 1 mM ATP in BRB80 was flowed in

(5 min adsorption time). Microtubules, antifade, and adenosine 5'-(β,γ-imido)triphosphate

(AMP-PNP, a non-hydrolyzable ATP analogue) in BRB80 buffer (128 nM total tubulin

concentration, 20 mM D-glucose, 0.02 mg/ml glucose oxidase, 8 µg/ml catalase, 1%

72

dithiotreitol) were perfused into the flowcell and allowed time (~10 min) to bind to the kinesin-

coated surface. Alignment solution (buffer, antifade, and 1 mM ATP, no microtubules) was

perfused into the flowcell to initiate motility. To achieve high flow rates for alignment, the

alignment solution was added continuously to one end of the flowcell while being removed from

the other end of the flowcell with a P200 pipette tip (Neptune esp, VWR, West Chester, PA)

connected to a vacuum line (Fisherbrand, ¼ inch inner-diameter, 3/32 wall width) connected to a

vacuum pump (0.02 HP, Barnant Company, Barrington, IL). Each alignment used

approximately 700 µl of alignment solution and the alignment time (between 75 s and 200 s)was

recorded. After the solution was used, a new solution containing buffer, antifade, and 1 mM

AMP-PNP was perfused into the flowcell. A solution of buffer, antifade, and 0.1%

glutaraldehyde was then perfused into the flowcell and left for 6 min. The glutaraldehyde

solution was exchanged with a buffer and antifade solution and the fixed array was then imaged

using a 40x Nikon S Fluor oil objective (NA 1.3, WD 0.22, Nikon, USA).

Imaging and Measurement

The flowcells were imaged using a Nikon Eclipse TE2000-U microscope (Nikon, Melville,

NY), an X-Cite Hg-arc lamp (EXFO, Ontario, Canada), a rhodamine filter cube (#48002,

Chroma Technologies, Rockingham, VT, a 40x (Nikon S Fluor, NA 1.3) or 100x (Nikon Plan

Apo, NA 1.45) oil objective, and an iXon EMCCD camera (Andor DV885-JCS-VP, South

Windsor, CT). The microtubule orientation was analyzed using ImageJ (v.1.41o, National

Institutes of Health, USA). The end-points of the microtubules were marked and this

information was used to calculate the orientation. Only straight microtubules were included in

the measurement of the orientation. No bias against long microtubules was introduced. The

73

flow rate was calculated by dividing the total volume of alignment solution perfused by the time

taken to pass through the flowcell.

The shear rate was calculated assuming a parabolic flow profile, a width of 5 mm, a

height of 100 µm, and an average volumetric flow of 5 µl/s. The volumetric flow rate was used

to calculate the maximum velocity, which in turn was used to calculate shear rate.

Figure 6-1: Concept of a macroscale hybrid linear motor using nanoscale stators and forcers. A)

Microtubules are aligned in an isopolar manner using fluid flow and fixed to the surface, creating a nanoscale stator. A macroscale kinesin-coated object placed onto the stator surface acts a nanoscale forcer. B) Multiple layers of stators and forcers can be layered to produce macroscale forces and displacements.

10 mm

A

B

20 n

m

Kinesin

Macroscale Object Motion 10 µm

Microtubule

74

Figure 6-2: Creating an isopolar microtubule array. A) Microtubules diffuse to the kinesin-coated surface and bind with random orientations in the presence of AMP-PNP. B) Microtubule motility is initiated with the introduction of ATP, the microtubules are aligned using fluid flow, and motility is stopped with reintroduction of AMP-PNP. C) Following alignment, the microtubules are cross-linked to the surface using glutaraldehyde, without visibly damaging the microtubules.

Figure 6-3: Analysis of isopolar microtubule arrays. A) An example of the microtubule orientation distribution before (blue) and after (red) alignment. B) Average orientational efficiency of microtubule arrays. Before and after alignment, the arrays have efficiencies of 68±30% and 94±13% (mean±std), respectively.

A B C

0

10

20

30

40

5 15 25 35 45 55 65 75 85

Num

ber o

f Mic

rotu

bule

s

Orientation (degrees)

0

20

40

60

80

100

Orie

ntat

iona

l Effi

cien

cy (%

)

Pre- Post-Alignment Alignment

A B

75

Figure 6-4: Force-generation of a microtubule array as a function of microtubule orientation. Arrays with high orientational efficiencies (0.8 < η < 1, shaded area) produce the greatest forces.

-1

-0.5

0

0.5

1

0 30 60 90 120 150 180

Nor

mal

ized

For

ce

Average Microtubule Orientation (degrees)

F = <cos(α)>

76

CHAPTER 7 CONCLUSION AND OUTLOOK

Bionanotechnology is a nascent field with many challenges. This dissertation focused on

four specific aspects of hybrid bionanodevice development. Firstly, a freely-programmable

patterning method, termed light-to-heat-conversion, was developed that can direct-write proteins

onto thermo-responsive polymer surfaces with 2 µm resolutions.239 Secondly, the temperature-

dependent activities of two species of motor proteins were characterized.240 Thirdly, cellular

control strategies were incorporated into a hybrid device and new insights into future control

strategies were found.241 Finally, a hybrid linear motor was designed and the microtubule-based

stator was optimized.242

Although the specific knowledge gained in the research performed has limited direct

applicability, many valuable general insights were made that are applicable across the field.

Firstly, many current protein patterning methods are adapted from photolithographic processes

developed for microprocessor fabrication. While these adaptations have proven useful,

mimicking the self-assembly and fabrication processes found within cells will likely prove more

powerful and versatile than the strictly top-down approach. Secondly, many proteins have not

been thoroughly characterized, and knowledge of the specific activity of the proteins used in

hybrid devices is a vital design parameter. Thirdly, hybrid devices utilizing chemical control

methods cannot effectively address individual components, nor is this necessarily the ideal

approach. A general approach to controlled activation of a hybrid kinein-microtubule system

was developed, but the same approach can be used for a variety of hybrid systems. Finally, the

actuator that will be produced based on the stator developed in the last chapter can be directly

incorporated into the next generation of hybrid devices that will be chemically, rather than

electrically, controlled. Furthermore, the solutions to the design and fabrication challenges

77

experienced can be applied to the development of other hybrid actuators, even if the kinesin-

microtubule system is not used.

Biological systems and components have been optimized for their in vivo functions, but

incorporating them into synthetic environments is a non-trivial task. Although the laws of

physics do not change at smaller size-scales, the forces that dominate interactions differ from the

macroscale and at times are counter-intuitive.243 Cells have evolved into versatile nano- and

microscale machines, and replicating their capabilities will produce great advances in the fields

of biotechnology and nanomedicine.

The potential of hybrid bionanodevices is nearly limitless. At their current and simplest

level, bionanodevices can duplicate the analytical capabilities of an entire laboratory with a

single device less than 1 mm in length.106 As techniques and technologies improve, device

dimensions can decrease and functionality can increase. Current hybrid devices are used in a

manner similar to lab-on-a-chip devices, where solutions with analytes are added to the external

bench-top device. By using active transport rather than microfluidic pumps, hybrid biosensors

can be decreased to microscale dimensions. This would allow the devices to be injected into

analyte-containing media (i.e. blood, water works, etc) and function autonomously, rather than

injecting sample fluids into the device, a slow and cumbersome process. Ultimately, hybrid

bionanodevices could function in any number of roles, but the key challenges will be developing

effective methods of controlling and communicating with the devices.

78

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BIOGRAPHICAL SKETCH

Rob Tucker was born in 1982 in Shelton, Washington. Despite never staying in any one

place for more than five years, Rob loves to travel, both within the United States and throughout

the world. In 2005 he earned a Bachelor of Science from the Department of Chemical

Engineering at the University of Washington-Seattle. Rob continued his education with his

undergraduate advisor at the University of Florida, Gainesville, where he earned his Ph.D. in

August 2009. Although he enjoyed his time in graduate school, Rob is excited to move into

industry, where he hopes to design, develop, and produce biomedical devices and point of care

diagnostic systems using hybrid bionanotechnologies.

© 2009 Robert Tucker - University of Florida

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