|UFDC Home||myUFDC Home | Help|
This item has the following downloads:
1 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
2 2009 Robert Tucker
3 To my Family and Friends for their unwavering support
4 ACKNOWLEDGMENTS First and foremost, I thank my advisor Dr. Henry Hess for his guidance, friendship, and support th roughout 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. Christ opher 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 n o t 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 Bruer, Gero Fink, Veikko Geyer, Dr. Leonid Ionov, Till Korten, Dr. Cecile Leduc, Doreen Naumburger, Bert Nitzsche, Cordula Reu ther, 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.
5 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 Meth ods ................................................................................................................ 46 Kinesin and M icrotubule P reparation ................................................................................. 46 Variable T emperature M otility A ssay ................................................................................ 46 Measurement of T emperature ............................................................................................. 47 Measurement of V elocity .................................................................................................... 47
6 5 CONTROL: ADAPTING CELLULAR CONTROL STRATEGIES TO HYBRID BIONANODEVICES ................................................................................................................. 50 Introduction ................................................................................................................................. 50 Results .......................................................................................................................................... 51 Discussion .................................................................................................................................... 55 Materials and Methods ................................................................................................................ 58 Kinesin and Microtubules ................................................................................................... 58 Caged -ATP and Hexokinase ............................................................................................... 58 Moti lity Assays and Microscopy ........................................................................................ 58 Velocity and Radius M easurements ................................................................................... 60 Numerical M odel ................................................................................................................. 60 Analytical Model ................................................................................................................. 62 6 ACTUATION: ASSEMBLY OF AN ISOPOLAR MICROTUBULE ARRAY TO PRODUCE MACROSCALE FORCES USING NANOSCALE COMPONENTS ............... 67 Introduction ................................................................................................................................. 67 Results .......................................................................................................................................... 68 Discussion .................................................................................................................................... 70 Materials and Methods ................................................................................................................ 71 Kinesin and Microtubules ................................................................................................... 71 Micr otubule Array Preparation ........................................................................................... 71 Imaging and Measurement .................................................................................................. 72 7 CO NCLUSION AND OUTLOOK ............................................................................................ 76 LIST OF REFERENCES ................................................................................................................... 78 BIOGRAPHICAL SKETCH ............................................................................................................. 91
7 LIST OF FIGURES Figure page 1 1 The State of the Art of Artificial Organs .............................................................................. 17 1 2 A hybrid bionanodevice ......................................................................................................... 18 2 1 Mi crotubules 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 Rela tionships 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
8 6 3 Analysis of isopolar microtubule arrays ............................................................................... 74 6 4 Force -generation of a microtubule array as a function of microtubule orientation. .......... 75
9 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 nex t 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 t he nanoscale, new fabrication methods must be developed. A new rapid prototyping technique using confocal microscopy was used to produce freelyprogrammable highresolution 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 poten tial into
10 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 de signed which consists of a molecular motor -coated synthetic macroscale forcer and a microtubule -based stator. M ethod s to create and characterize the stator w ere developed, which can be used to optimize the force generation of the device
11 CHAPTER 1 INTRODUCTION Within almost every cell exists complex nanoscale machinery, capable of repairing damage,1, 2 communicating ,3, 4 and creati ng 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 i mmune cells operate autonomously, moving freely throughout the body to identify foreign objects and pathogens .10 12 Other cell s cooperate to better suit their tasks, working in large groups to form tissues and organs.13 The human heart is one of the few organ s to be successfully replaced with a permanent ly implant ed artificial device. Totally implantable artificial hearts were recently approved by the United States Food and Drug Administration, but have limited life time s and duplicate only the mechanical aspects of natural hearts .1417 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 t he heart has be en 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, 2126 Although the heart as a whole responds to these signals, it is the individual card iomyocytes -heart muscle cells comprising the heart that detec t 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 t he pancreas has been studied in great detail but no permanently implantable artificial device to duplicate its functions exist s The pancreas fulfills two roles within the body: as part of the exocrine system it aids in digestion and as part of the endocrine syste m it regulates
12 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 chr onic autoimmune disease wherein the body can no longer regulate blood glucose levels This disease affect s over 18 million people in the United States alone .27 Despite extensi ve research, no cure for diabetes exists, and the current treatments are crude. D iabetes 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 ha s been mimicked with multiple macroscale devices (Fig. 1 1 B), the cell level and exocrine functions have not Implantable glucose sensor s and insulin pumps will significantly reduce the impact of diabetes on patients lives, but will not replace a pancreas. For this, a synthetic biomedical device capable of replicating cell level func tions 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 T he 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 Cell s 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 .3537 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 ta rget location.41, 42 In vivo t he mode of m aterial transport depends on the type and size of the material being transported .43 Larger molecules, proteins,
13 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 indivdual ly or in group s 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 te chnologies 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 functional ity 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. Finall y, current technologies have almost no ability to individually address nanoscale objects, a feat cells perform continuously, i.e. proteins, vesicles, and filaments. First ly p roteins are extremely sensitive to environmental conditions including tempera ture, ionic concentration, pH and radiation .4851 In contrast to the wide range of environmental conditions used during fabrication of synthetic devices such as insulin pumps and electronics,
14 fabrication of hybrid bionanodevices requi re new methods that are significantly closer to physiological conditions However, a s device size decrease s new combinations of fabrication techniques can be utilized which take advantage of the bottom up self assembly of biological systems and the top -d own 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.5255 A rapid -prototyping method, termed light to -heat conversio n (LHC), was developed and used to pattern functional proteins onto thermoresponsive 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 contra st to photolithography that uses a fixed mask to pattern, the direct -write LHC procedure is freely programmable. Patterns with 2 m resolutions were written and smaller resolutions are possible. Secondly, b iological systems respond strongly to temperature changes. Warm -blooded animals use significant amounts of energy to maintain their body temperature, which allows them to have compara tively simpler proteins that operate within a small temperature range .56 Small organisms and c old -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 implement ed 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 man y 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 thermodynamical ly
15 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 temperatu re range.49 The activity of this kinesin was compared with the act ivity of conventional kinesin derived from drosophila melanogaster in an attempt to develop a temperature insensitive bionanodevices. Th i rdly, h ybrid bionanosystems direct ly conver t chemical energy into mechanical movement in contrast to many current bi omedical 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, e lectrically 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, m acro and microscale actuators have been produced primarily using a topdown approach, although combinations of bottom up self assembly and topdown approaches have been previously developed.60 In n ature, 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 kilone wtons. T he force generat ion by single or small groups of molecular motors ha s been investigated in vitro in great detail, but biomimetic actuators based on large arrays of motor
16 proteins ha ve 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 c an no t be achieved without the combination of topdown and bottom up approaches. Hybrid Bionanodevice s The intracellular transport system found within cells can be mimicked, augmented, and integrated into artificial bionanodevices. Hybrid devices composed o f 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 det ect 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 captur e 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 illum inated 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.
17 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. Artific i al hearts are macroscale pumps that cannot replicate the intercel lular 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 oper ate at the macroscale. Adapted from 68 71. A B
18 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 specifi c to the analyte and fluorescently tagged. The tagged particles are transported to the collecting chamber where the signal is detected. Adapted from 72.
19 CHAPTER 2 BACKGROUND AND EXPERIMENTAL TECHNIQ UES 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 Th e 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 an d microtubules.73 Actin, with its compl e mentary motor myosin i s 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 ce ll. 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 a re located in the nuclear membrane to maintain the stability of the nucleus,74, 75 vimentin is foun d 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 filam ent 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 pa rt structural hierarchy beginning with heterodimeric
20 and 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 microtu bule can vary between 11 and 17.8185 Microtubules were first discovered in the 1950s and considered static struc tures,86, 87 but their continuous in vivo polymerization and depolymerization termed dynamic instability was disc over ed 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 d oes. 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 -frame s, 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, m icrotubules and actin filaments are us ed as roadway s 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 moveme nt (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
21 the -tubulin of the microtubule, a nec k 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 -tubulin. The binding of the lead ing head to the microtubule causes the rear adenosine di -phosphate ( ADP ) bound head to swing forward 16 nm to the next tubulin. ATP -hydrolysis in the rear head releases ~20 kBT66, 91 of energy and form s 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 10001, 7 00 nm/s under physiological con ditions .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 micr otubules 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 3 A ). The motility assay is performed as follows: First, a f lowcell 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 ( = 535
22 nm), produces photons ( 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 -glucono1,4 -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 E pi 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 1994.98102 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 l ight.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 < < 800 nm). Selection of the filter set depends on the fluorescent dyes being us ed. Regardless of the specific fluorophores being used, the set consists of three filters known as excitation, dichroic, and emission filters The excitation
23 filter is a band pass filter that passes a small subset of the full spectrum ( excitation 25 nm) produced by the arc lamp exc iting 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 th e sample. The dyes within the sample absorb the excitation photons and release lower energy, longe r wavelength photons ( emission) Th e 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 pass es only light originating from the dye in the sample ( emission 35 nm) while blocking all other wavelengths of light Different filter sets can be used in conjunction with different fluorescent dyes, which a llows 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. T he 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 ( ), 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) a re 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 in dex 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,
24 is the wavelength of light used to illuminate the sample, and NA is the numerical aperture of the lens r (21) 2NA 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 highintensity laser, in c ontrast 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 a nd 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, t he fluorescent dyes in the illuminated region of the sample are excited and emit lo wer 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 c onfocal 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 rat ios 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 produ ces 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, i maging the sample with a highintensity laser allows for shorter illumination periods that produce the same number of photons from the sample, resulting in higher signal to noise ratio s 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, t he 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 i s 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 produc ing 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 doe s have two distinct disadvantages that can limit it s use. A s 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. Howe ver, i n 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. Second ly 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 microscop y 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 hetero dimeric proteins known as (light brown) and -tubulin (dark brown), which bind G T P and G D P. 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 stru cture. 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 tubulin of the microtubule heterodimers. B) The binding of the leading ATP bound head causes the rear ADP -bound head to swi ng forward 8 nm to the next 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 m. Adapted from 105.
28 Figure 2 3: I n 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 cover slip 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 m icrotubule towards the minus -end. (C) The fluorescently labeled microtubules are imaged using epi -fluoresence microscopy and a CCD camera A dapted from 106. A B C + end Casein Microtubule movement Objective Slide Spacers C overslip Flow in Flow out
29 Figure 2 4 : Diagram of an e pi -f luorescence m icroscope and s pectr a of a f ilter c ube (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. A Sample Objective Camera Arclamp Excitation filter Emission filter Dichroic filter HQ535/50x Q565 LP HQ 610/75m Transmission (%) 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 ( ). 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* sin Objective lens Oil Air n=1.5 n=1.0 Slide Media Coverslip Imaging Plane
31 Figure 2 6 : Confocal Microscopy Schematic. Confocal microscopy is a type of fluorescence microscopy, but higher spati al and temporal resolution image s 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 BIOFU NCTIONAL PROTEINS ON THERMORE SPONSIVE SURFACES Introduction Patterning functional proteins onto artificial substrates is of interest in the development of n anotechnology, tissue engineering, biosensors, and cell biology.107119 Towards this end, a number of chemical patterning metho ds based on optical lithography,120122 atomic force microscopy (AFM) ,123, 124 printing techniques ,125 chemical vapor deposition (CVD)126128 have been recently developed. While each of these m ethods 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 l ow 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 co mbined 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 ( ~ 350 nm) of ten 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 photogenerati on 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 thermores ponsive polymer surface capable of binding proteins and maintaining their functionality. Previously it was show n that poly(N -isopropylacrylamide) (PNIPAM) could be used to control the binding of proteins onto thermoresponsive surfa ces.131 132 The re, the conformation of PNIPAM molecules in aqueous solution was switched in a spatially unstructured manner between the collapsed state at T > 33C (protein-binding conformation) and the swollen state at T < 30C (protein -repelling conformation). Here, the approach was extended by using optical signals to generate heat in a highlylocalized manner (Figure 3 1). T o efficiently convert light to heat, (i) solution -bound malachite green molecules133 (Figure 3 1 A ) or (ii) substrate -bound black -ink layers (Figure 3 1 B) 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 fur ther protein binding. The swollen polymer chains also protected the patterned proteins during consequent fluid exchanges, providing a means to sequentially pattern multiple protein s pecies on the same surface without the need for specific linker molecules or elaborate surface preparation. Preparation of PNIPAM S urfaces PNIPAM layers were prepared via a two -step procedure similar to methods described previously131 on glass coverslips (for solution-b ased 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. Highgrade mica substrates (Ted Pella, Inc.) were dip -coated with edding T100 black ink (edding
34 International GmbH ), dried at 120C for 1 h and baked at 320C 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, M n = 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 pol y( N isopropylacrylamide) (PNIPAM COOH, M n = 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 180C. 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 thi cknesses 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 ( = 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 P hotopatterning of P roteins To demonstrate the two LHC approa ches, 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 f irst 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 c onfocal microscope (Zeiss LSM 510, Carl Zeiss GmbH, Germany) using a 99 mW UV laser ( = 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 s/pixel dwe ll 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 1 C). Using this method, a ~2 m feature size was achieved as seen by the line widths shown in Figure 3 2. As the patterning method wa s based on confocal microscopy, any pattern size or shape with a resolution limit of 2 m 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 glo bular 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 surfa ce coated with immobilized PNIPAM, in place of the bottom coverslip (see Figure 3 -
36 1 D ). The nonpatterned 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 combin ation 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 G liding M otility A ssays To test the efficacy of the LHC layer, gliding motility assays were p erformed directly on the blackene d 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 inv erted in the gliding assay, and microtubules 5 10 m in length gliding over substrate -bound kinesin Because the gliding speed of microtubules on kinesin surfaces wa s known to strongly depend on temperature it was possible to estimate the solution tempera ture when illuminated .140 Motility experiments were performed in a 5 mm wide flowcells constructed from a glass cover sli p (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 L of a kinesin solution containing 2 g/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 7C 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 micr otubule gliding velocities were v = 0.19 0.04 m/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 m/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 6C to approximately 22C. This experiment provides reasonable evidence for conversion of light into heat on the LHC surface. Testing the F unctionality of P atterned P roteins 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 35C for 5 min. The flowcell was then cool ed down to room temperature (25 C) and the kinesin solution (10 g/mL) was perfused in. A section of the blackened substr ate surface was then illuminated through a 100x objective (Zeiss, Plan Neofluor, NA1.3, oil) using g reen light ( = 480 nm, HBO 100 (Osram) arc lamp for 10 s). After the illumination was switc hed off, microtubule -containing motility solution was perfused i nto the flowcell at room temperature and the entire flowcell was heated to 35C 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 (35C) 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 fi lms 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 o ne 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 resolution s should be attain able 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 fa brication 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 ( = 360 nm) laser beam irradiation usin g 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 usi ng UV ( = 351 & 364 nm) and green ( = 633 nm) light. The lines are approximately 2 m 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 ( = 480 nm). After the patterning, the kinesin retained its biological functionality and provided continuous gliding of microtubules over the patterned surface at high temperature (35C). Cooling down to room temperature (25C) led to the release of microtu bules 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 O F PROTEIN ACTIVITY F OR THE DEVELOPMENT OF TEMPERATURE INSENSITIVE HYBRID D EVICES 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 10C increase in temp erature at saturating substrate concentrations.143 Traditionally, temperature is closely contro lled 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 i nternal 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, metaboli c 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]
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 structur es,152160 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 electroosmotic flow.169 Bhm et al.170 previously reported that the Km values for porcine kinesin 1 at temperatures of 25C and 35C are 6610 M and 798 M, resp ectively. Kawaguchi et al. determined that the activation energy of bovine kinesin1 is 50 kJ / mol, implying a Q10 at saturating ATP concentrations equal to two.171 This suggests that the activity change per 10C change in temperature for small ATP concentrations is significantly smaller than two, implying that the increasing Km partially compensa tes for the increasing vmax. Similarly, ATP consumption assays for Thermomyces lanuginosis kinesin 3 determined an activation energy of 949 kJ / mol, and an increase in the Km from 427 M at 25C to 1.60.3 mM at 50C.49 At low ATP concentrations the Q10 is 0.7 and thus the Thermomyces kinesin activity falls with increasing temperature despi te 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 19C to 34C 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 V elocity measurements for microtubules gliding on Drosophila kinesin 1 (Figure 4 1) show ed 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 o f 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 34C data point of the Thermomyces data was not included in the fit of vmax and Km since the reduced vmax indicate d part ial 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 535.1 kJ / mol (Q10=2.04) and 383.3 kJ / mol (Q10=1.67) for Drosophila kinesin 1 and Thermomyces kinesin3, respectively. Independent of temperature, the Km values were found to be 7010 M and 22090 M for Drosophila kinesin1 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 wa s not surprising that the measurements in the temperature interval from 19C to 34C 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 wa s 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 45C.49 A second preparation of Thermomyces kinesin 3 showed significantly altered motility (more interruptions of gliding motion and more stuck mic rotubules) and a Km of 175 M at 23C, which indicate d that variations between preparations exist ed It wa s more challenging to reconcile the increase of Km for porcine kinesin -1 observed by Bhm et al.170 with the observations of the Thermomyces and Drosophila kinesin A possibl e explanation for Bhm 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 kinesins 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 conclude d that stabilization against temperature changes will require the design of a suitable enzymatic network, adding to the complex ity of the device similar to the recently demonstrated enhanced robustness of a DNA nanomotor.176
46 Materials and Methods All chemicals we re 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 37C 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 M paclitaxel. Variable temperature motility assay A new flowcell was assembled for each ATP concentration tested, which consisted of a square coverslip (2222 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 g/mL microtubules, casein (0.2 mg/mL), an an tifade solution (20 mM D rad Laboratories, Hercules, CA)), an ATP regeneration system (2000 Units/L creatine phosphokinase, 2 mM creatine phosphate ref. 174) and ATP (10 M, 25 M, 50 M, 100 M, 200 M or 1000 M) was introduced, and the edge s 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 concentra tions tested were 10 M, 100 M, 200 M, 1000 M, and 5000 M. 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 (19C, 23C, 26C, 31C, or 34C) a nd automatically regulated by a temperature controller (Warner Instruments Inc., TC 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 i nterest 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 flowcell s. 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=19C) and the flowcell was heated to each new temperatur e 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 temperat ure for various ATP concentrations. Black stars are the data published by Kawaguchi and Ishiwata171 for 1 mM ATP. (B) Microtubule gliding velocity on Thermomyces kinesin3 as function of temperature for various ATP concentrations. 20 24 28 32 0 500 1000 1500 2000 2500 1000 M Kawaguchi 1000 M 200 M 100 M 50 M 25 M 10 MVelocity (nm/s)Temperature (oC)20 24 28 32 5000 M 1000 M 200 M 100 M 10 MA 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. Dro sophila kinesin 1 full squares, blue; Thermomyces kinesin 3 open triangles, red. 0 50 100 150 200 1000 0 250 500 750 1000 1250 1500 19 C 23 C 26 C 31 CVelocity (nm/s)[ATP] ( M) 34 C 0 100 200 2500 5000 0 500 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) vmax (nm/s)Temperature ( C)vmax = 8nm x 1.1x109s-1 x e-4600K/(T( C)+273K)18 20 22 24 26 28 30 32 34 0 100 200 300 Km = 70 M + (0.1 0.7) x (T( C)-25 C) Km ( M)A BKm = 218 M (0.3 0.3) x (T( C)-25 C)
50 CHAPTER 5 CONTROL : ADAPTING CELLULAR CONTROL STRATEGIES T O HYBRID BIONANODEVICES Introduction A challenge for nanotechnology is the dynamic and specific control of nanomachines by the user. Molecular shuttles, consisting of cargobinding microtubules propelled by surface immobilized kinesin motor proteins, are an example of a nanoscale system which ideally can be selectively activated at programmable locat ions and times. Here a biomimetic solution is discussed where activating molecules we re delivered locally via photolysis of a caged compound and subsequently sequestered in an enzymatic network. The controlled sequestration of the activator not only creat e d a rapid deactivation when the stimulus wa s removed, but also sharpen ed the concentration profile of the rapidly diffusing activator. This improvement c a me at the expense of a reduced efficiency in the utilization of the activator molecules, suggesting t hat these nanosystems are most efficiently addressed as swarm and not as individuals. This work represent ed 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 c ontrol 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 seque stering 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 acti vation an integral part of the design as in the light -driven synthetic motors based on rotaxanes or catenanes ,182 o r 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 patte rned 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 th e 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, ar o se from the rapid diffusion of ATP (D=3x1010 m2/s) ,187 which outpace d 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 limit ed the average distance an ATP molecule c ould 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 m or 25 m lead to a dramatic improvement in
52 the temporal control over the shuttle activation. The shuttles immediately (<1s) respond ed to the illumination by beginning to move. After the illumination end ed the shuttle velocity dropped by 1/e on a timescale of 10 s without sequestration by hexokinase to 1 s with hexokinase p resent Previously,162 uniform illumination of the cell led to deactivation on a timescale of hours to minutes in the absen ce and presence of hexokinase, respectively. Of course, this improved temporal control wa s 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 25 0 ms pulses cause d the shuttles to move a distance equal to the distance traveled with a single pulse equal to the total duration of illumination, which implie d that the average translation per pulse c ould be tuned to less than 50 nm. While the diffusion of the activating molecule, ATP, benefit ed the control of activation in the temporal domain, the situation wa s reversed in the spatial domain. The velocity of individual shuttles as function of their distance from the center of illumination (Fig. 5 2 A ) t o determine velocity profiles was measured Since the velocity profiles of pulsed and continuous illumination were very similar the discussion focus es on the steady state between the release, diffusion and sequestration of ATP reached during extended illu mination. 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) show ed an approximately exponential decay of the velocity with increasing radius. As the hexokinase concentration in the solution wa s 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 c oncentration profile by a factor of 5 7. Changing the
53 radius of the illumination zone from 25 m to 15 m reduce d the observed velocities to a third, roughly in proportion to the area of the illumination zone (at constant light intensity). To model th e 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 numeri cally 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 M 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 aft er 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 wa s 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 prove d 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 we re 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 m2/s wa s the diffusivity of ATP, Km=0.12 mM wa s the Michaelis constant of hexokinase with respect to ATP, and A wa s the activity of the hexokinase solution. The approximate solution191 of t he 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 Equatio n 51 where C0 is the ATP concentration at the boundary of the illuminated zone. 0 *exp (5-1)i irr r CC r r Velocity profiles (Fig. 5 3) we re calculated from Equation 5 1 using v=fvmaxC/Km kin, where f wa s the above mentioned factor accounting for slowing motor interactions, Km kin wa s the Michaelis constant for ATP consumption by kinesin, and small ATP concentrations we re assumed. The experimental data we re fit to the above expression for v outside the illumination zone while the velocity within the zone wa s assumed to be constant and equal to v0=fvmaxC0/Km kin. The fit to the six experimental profiles utilize d six values of C0 as free parameters while the value of r* wa s obtained from the known values of Km, D and A (r* = 210 m, 66 m and 21 m for 50, 500 and 5000 units/L of hexokina se, respectively), and the values of f we re the same as in the numerical simulations. The values of C0 obtained from the fits we re 1.4 M, 0.9 M, 0.5 M for ri = 15 m at 50, 500 and 5000 units/L hexokinase activity and 3.3 M, 1.9 M, 0.7 M for ri = 25 m 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 c ould be determined. At low light intensities, the ATP generation wa s not limited by the diffusive influx of caged ATP into the illumination region, but wa s given by G =
55 CcATPkI, where CcATP wa s the constant concentration of caged ATP, k wa s the ATP uncaging rate constant and I wa s the UV light intensity .192 This result ed in the concentration formula shown in Equation 5 2. 0 2 (5-2) 11cATPhv iCkI C D r r Based on the known values for CcATP, D, r*, ri, k and the fit values for C0, this implie d that the average illuminating UV i ntensity is 2 mW/cm2 for both pinhole sizes. The analytical solution illustrate d the opposing trends in achieving localized control: A reduction in the characteristic ATP diffusion distance r* (increase in hexokinase activity) le d to reduced ATP levels (F ig. 5 4A) but at the same time narrow ed the ATP plume as measured by the radius at which the shuttle velocity was reduced to 10% of the maximum (r10%, Fig. 5 4 B). Interestingly, if the absolute concentration gradient C0/r10% wa s used as the performance metric of the control process (Fig. 5 4C), it bec a me apparent that an optimum for the characteristic diffusion distance existed Since r10%i+r* for r*<10ri (Fig. 5 4B), by finding the maximum of C0/(ri+r*) the optimal value of the characteristic diffusion distance r* c ould be determined to be approximately equal to twice the illumination radius ri (Fig. 5 4C). Discussion This suggest ed a strategy to achieve a desired level of control in the case of cylindrical symmetry: (1) Def ine 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 illum ination 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 le d to a line ar decrease of the generated gradient and a quadratic decrease in fuel efficiency, which wa s defined here as the ratio of the number ATP molecules used by motors to the total number of ATP molecules released. The decreasing gradient c ould be compensated by increasing the intensity of the light source, but the amount of ATP which c ould be released wa s ultimately limited by the influx of caged ATP into the illumination zone. However, the light intensity at this limit wa s expected to lead to unacceptable level s of photodamage. The above described experiments, simulations and calculations demonstrate d 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 improve d the spatial and temporal control, even a restriction of the activation zone to tens of micrometers reduce d the activation level drastically. At the same time the vast majority of the released control molecules d id no t 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 li mitations, 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 switchi ng 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.203208 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 macros cale. 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 Hofstadters anteater, you must not take an ant for the colony .210 The engineering task is then to tailor the swarm behavior, e.g. by limi ting dispersion169 or by self -generating ATP157 rather than the individual nanotransporter.
58 The insights generated from this investigation of control via diffusi ng 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 Materials and Methods Kinesin and Microtubules Full length, wild -type kinesin from Drosophila melanogaster expresse d 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 m M EGTA, 5 mM MgCl2, 1 mM guanosine 5 triphosphate, and 5% dimethylsulfoxide) was added to a 20 g aliquot of rhodamine labeled tubulin (TL331M, Cytoskeleton, Denver, CO) resulting in a final tubulin concentration of 32 M. The solution was kept at 37C f or 30 minutes and then diluted 100-fold in BRB80 (80 mM PIPES, 1 mM MgCl, 1 mM EGTA, pH 6.9) containing 10 M taxol for stabilization and kept at room temperature (20C). Caged ATP and Hexokinase Photocleavable 1 (4,5 Dimethoxy2 -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 1000400,000 Units/L. These stock solutions w ere then diluted in motility solutions resulting to final concentrations of 25 5000 Units/L. The KM for hexokinase with respect to ATP is 120 M. Motility Assays and Microscopy The motility assays were performed in 100 m 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 containi ng the caged ATP was flowed into the flowcell (32 nM microtubules, 500 M caged -ATP, and an oxygen -scavenging system consisting of 20 mM D -glucose, 0.02 mg/ml glucose oxidase, 8 g/ml catalase, 1% dithiotreitol in BRB80). D -glucose is simultaneously the su bstrate 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 wa s 120 M, the hexokinase activity wa s 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 rho damine 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 m (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 UV340 Light Meter, Ambient Weather, Tempe, AZ). Within the flowcell, this light power of 0.8 W and 1.3 W was focused into a near cylindrical cone with a radius of 15 and 25 m
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 resolve d All assays were performed at 20C. 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 ac tivities 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. 52). The velocity wa s 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 c oncentration 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 differe ntial 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 22 22 m-[ATP][ATP]D[ATP] [ATP] D G(r,t) A (5-3) t rr r K [ATP] Here, the ATP concentration depend ed on th e 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 wa s 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 m 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 s hown in Equation 5 4. The Km for hexokinase was set to 120 M. The resulting ATP concentration profile was then used to calculate the molecular shuttle velocity assuming Mic haelis -Menten kinetics of kinesin with a vmax of 1000 nm/s, a Kmkin of 25 M, inhibition constants (Ki) for ADP and caged-ATP of 35 M and 200 M and is shown in Equation 5 4. max kin kin kin mm m caged-ATP ADP iiV[ATP] V f (5-4) KK [ATP] K [caged-ATP] [ADP] KK 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 65100, 60, 40, 20, 6, 1 nm/s, respectively. The time to reach s teady -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 wa s constant, and that the consumption of ATP by kinesin compared to hexokinase wa s negli gi ble. The steady state diffusion and sequestration of ATP ou t side the illumination zone (ass uming cylindrical symmetry) is shown in Equation 55. 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 21 0 (5-5)mCCA DC rrK r Equation 5 5 can be rewritten as shown in Equation 5 6 where r*2 = Km/DA 22 2 220 (5-6) CCr rrC rrr Equation 5 6 is the modified Bessel equation for which one of the solutions is shown in Equation 5 7 : 191 0 (5-7) r CaK r 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. 0 0 0 (5-8) *iC r CK r r K r 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) *iirrr CC rr
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) we re 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 we re measured as a function of the distance from the center of illumination for illumination zones with a radius of 25 m and 15 m 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 50 100 150 200 0 1 0 Units/L 50 Units/L 500 Units/L Speed (nm/s)Radius (m ) 0 5 5000 Units/L15 m Radius of Illumination 5000 Units/L25 m Radius of Illumination0 50 100 150 0 Units/L 50 Units/L 500 Units/L
66 Figure 5 4: Relationships between the maximum ATP concentration (A), the distance at which the AT P 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 r adius 15 m and 25 m.
67 CHAPTER 6 ACTUATION : ASSEMBLY OF AN IS OPOLAR MICROTUBULE A RRAY TO PRODUCE MACROSCALE FORCES USING NANOSCALE COMPON ENTS 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.214217 Motor proteins such as kinesin and myosin have been studied in great detail, both in vivo and in vitro .62, 65, 218223 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 ar rays of myosin, as seen in muscle.29, 30 Kinesin and myosin have been use d 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, le ngth of the motor unit (sarcomere), the density of
68 motors, and the orientational order of the components. From these design characteristics, it wa s learn ed what parameters are important in the design of a hybrid linear motor. An engineered microtubule ar ray 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 acti vity. 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 M icrotubules we re bound to a kinesin -coated s urface aligned with fluid flow to produce an isopolar array, and subsequently fixed to the surface using glutaraldehyde. The microtubules land with random orientations, and t he flexibility of the kinesin motors allow microtubule s to move in any direction o ver 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 1 00 m)229, 230 and the movement of the tip can be biased using low shear forces. Under flow, the lo west 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 is opolar microtubule array. The limitations of this alignment method are: 1) the flow rate must be high enough to reorient the microtubule tip s 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 con centrations, the ability of kinesin to bind to the microtubule lattice is impacted.231 The glutaraldehyde concentration was chosen to securely fix the microtub ules 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 w ith random orientation upon landing (Figure 6 2A), after alignment (Figure 6 2B), and after fixation with glutaraldehyde (Figure 6 2C). Th is project focuse d on the fabrication and characterization of the microtubule array. Microtubule arrays were deve loped previously by several research grou ps, b ut no means of quantitatively compar ing the arrays exist s .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 I n essence the efficiency is a measure of the average orientation of the microtubules within the microtubule array compared to an array with perf ect alignment. In Equation 6 1, orientation is the orientational efficiency, N is the number of microtubules in the array, and is the angle of the microtubule relative to the direction of fluid flow ( =0 ) in Figure 6 2. orientation 11 coscos (6-1)NN
70 Discussion An example of the orientational distribution before and after alignment is shown in Figure 6 3 A. 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 orientation al distribution had an approximately exponential decay. The average orientational efficiency of four arrays before (68 30% mean st an d dev) and after (94 13%) alignment is shown in Figure 6 3B. As expected, b oth 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 l/s although variatio n 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 w ithin the array is shown in Figure 6 4. Within the non-linear region of the plot ( 0.8 < < 1.0 ), only modest gains in the force generation capabilities of the array are made when increasing the orientational efficiency. Therefore, the optimum orientatio nal 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 m icrotubule 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 ( ) is a function of the orientational efficiency ( ), the microtubule surface density ( ), and the average length of the microtubules comprising the array (< L >). = L ( 6-2)
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. Materials and Methods Kinesin and Microtubules Full length, wild -type kin esin 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 g aliquot of rhodamine labeled tubulin (TL331M, Cytoskeleton, Denver, CO) resulting in a final tubulin concentration of 32 M. The solution was kept at 37C for 30 minutes and then diluted 100-fold in BRB80 (80 mM PIPES, 1 mM MgCl, 1 mM EGTA, pH 6.9) containing 10 M taxol for stabilization and kept at room temperature (20C). Microtubule Array Preparation M icrotubules 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' ( 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 g/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 flowcel l 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 P 2 00 pipette tip (Neptune esp, VWR, West Chester, PA) connected to a vacuum line ( Fisherbrand, inch inner -diameter, 3/32 wall width ) connected to a vacuum pump ( 0.02 HP, Barnant Company, Barrington, IL ). Each alignment used approximately 700 l of alignment solution and the alignment time (between 75 s and 200 s) was recorded. After the solution was used, a new s olution containing buffer, anti fade, and 1 mM AMP -PNP was perfused into the flowcell. A solution of buffer, anti fade, and 0.1% glutaraldehyde was then perfused into the flowcell and left for 6 min. The glutaraldehyde solution was exchange d with a buffer and anti fade 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 Ecl ipse TE2000U 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 c amera (Andor DV885JCS -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 orien tation. 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 m, and an average volumetric flow of 5 l/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 nm Kinesin Macroscale Object Motion 10 m 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 6830% and 9413% (meanstd), respecti vely. A B C 0 10 20 30 40 5 15 25 35 45 55 65 75 85 Number of Microtubules Orientation (degrees) 0 20 40 60 80 100 Orientational Efficiency (%) 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 < < 1, shaded area ) produce the greatest forces. 1 0.5 0 0.5 1 0 30 60 90 120 150 180 Normalized Force Average Microtubule Orientation (degrees) F = < cos ( )>
76 CHAPTER 7 CONCLUSION AND OUTLOOK Bionanotechnology is a nascent field w ith many challenges T his dissertation focused on four specific aspects of hybrid bionanodevice development First ly a freely -programmable patterning method, termed light to -heat -conversion, was develo ped that can direct -write proteins onto thermo responsive polymer surfaces with 2 m 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 specif ic knowledge gained in the research performed has limited direct applicability, many valuable general insights were made that are applicable across the field. First ly many current protein patterning methods are adapted from photolithographic processes de veloped 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 c omponents, 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 chal lenges
77 experienced can be applied to the development of other hybrid actuators even if the kinesinmicrotubule 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 capabiliti es of an entire laboratory with a single device less than 1 mm in length.106 As techniqu es and techn ologies improve, d evice dimensions can decrease and functionality can increase. Current hybrid devices are used in a manner similar to lab -ona chip devices, where solutions with analytes are added to the external bench top device. By using active transp ort 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 samp le 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 LIST OF REFERENCES 1. Watanabe, K.; Iwabuchi, K.; Sun, J.; Tsuji, Y.; Tani, T.; Tokunaga, K.; Date, T.; Hashimoto, M.; Yamaizumi, M.; Tateishi, S. Nucleic Acids Research 2009, X, (X), 1 18. 2. Turchi, L.; Fareh, M.; Aberdam, E.; Kitajima, S.; Simpson, F.; Wicking, C.; Aberdam, D.; Virolle, T. In ATF3 and p15PAF are novel gatekeepers of genomic integrity upon UV stress, 2008; PergamonElsevier Science Ltd: 2008; pp 92 92. 3. Hofmann, K. DNA Repair In Press, Corrected Proof. 4. Dorn, G. W. Journal of Molecular Medicine 2009, 88, (1), 19. 5. Inoue, S.; Salmon, E. D. Molecular Biology of the Cell 1995, 6, (12), 16191640. 6. Sleeman, J. E. Philosophical Transactions of the Royal Society of London. S eries A: Mathematical, Physical and Engineering Sciences 2004, 362, (1825), 2775-2793. 7. Kim, S. -K.; Welsh, R. M. Journal of Immunology 2004, 172, (5), 31393150. 8. Spellman, P. T.; Sherlock, G.; Zhang, M. Q.; Iyer, V. R.; Anders, K.; Eisen, M. B.; Brown P. O.; Botstein, D.; Futcher, B. Molecular Biology of the Cell 1998, 9, (12), 32733297. 9. Snyder, G. K.; Sheafor, B. A. American Zoologist 1999, 39, (2), 189198. 10. Zen, K.; Parkos, C. A. Current Opinion in Cell Biology 2003, 15, (5), 557564. 11. Ma y, R. C.; Machesky, L. M. Journal of Cell Science 2001, 114, (6), 10611077. 12. Guermonprez, P.; Valladeau, J.; Zitvogel, L.; Thery, C.; Amigorena, S. Annual Review of Immunology 2002, 20, (1), 621667. 13. Weber, K. T. Cardiovascular Research 2004, 64, ( 3), 381383. 14. FDA, U nited States Food and Drug Administration 2006. 15. Dowling, R. D.; Gray, L. A.; Etoch, S. W.; Laks, H.; Marelli, D.; Samuels, L.; Entwistle, J.; Couper, G.; Vlahakes, G. J.; Frazier, O. H. Journal of Thoracic and Cardiovascular Surgery 2004, 127, (1), 131141. 16. Gemmato, C. J.; Forrester, M. D.; Myers, T. J.; Frazier, O. H.; Cooley, D. A. Texas Heart Institute Journal 2005, 32, (2), 168177. 17. Kirklin, J. K. H., William L. Current Opinion in Cardiology 2006, 21, (2), 120126. 18. Houston, G. W.; Amy, L. T.; Alexandrina, U.; Xinwei, S. Reports on Progress in Physics 2005, (3), 545.
79 19. Sharaov, V. G., Todor, Anastassia V., and Sabbah, Hani N. Heart Failure Reviews 2005, 10, (2), 13824147. 20. Catena, E., Milazzo, F. Echocardiography and cardiac assist devices. 2007, 55, (2), 247265. 21. John, W. Artificial Organs 2008, 32, (10), 757760. 22. de Bold, A. J.; Borenstein, H. B.; Veress, A. T.; Sonnenberg, H. Life Sciences 1981, 28, (1), 89 94. 23. Pina, I. L.; O'Connor, C. The Journal of the American Medical Association 2009, 301, (4), 432434. 24. Lopaschuk, G. D.; Kelly, D. P. Cardiovasc ular Res earch 2008, 79, (2), 205207. 25. Kirchberger, N. M.; Wulfsen, I.; Schwarz, J. R.; Bauer, C. K. Journal of Physiology London 2006, 571, (1), 2742. 26. Lamas, S.; Lowenstein, C. J.; Michel, T. Cardiovascular Research 2007, 75, (2), 207209. 27. CDC, The 2007 Annual Report of the OPTN and SRTR: Transplant Data 19972006 In 2007. 28. Magdalena Annersten, A. W. Worldviews on Evidence -based Nursing 2005, 2, (3), 122130. 29. Dickinson, R. B. Journal of Mathematical Biology 2009, 58, (12), 81103. 30. Dickinson, R. B.; Caro, L.; Purich, D. L. Biophysical Journal 2004, 87, (4), 28382854. 31. Phng, L. K.; Gerhardt, H. Developmental Cell 2009, 16, (2), 196208. 32. Trautmann, A. Science Signaling 2009, 2, (56), 13. 33. Hill, D. B.; Plaza, M. J.; Bonin, K.; Holzwarth, G. European Biophysics Journal 2004, 33, (7), 62332. 34. Hirokawa, N. Science 1998, 279, (5350), 519526. 35. Rafelski, S. M .; Theriot, J. A. Annual Review of Biochemistry 2004, 73, 209 239. 36. Fletcher, D. A.; Theriot, J. A. Physical Biology 2004, 1, (1 2), T1 T10. 37. Lauffenburger, D. A.; Horwitz, A. F. Cell 1996, 84, (3), 359369. 38. Francis, K.; Palsson, B. O. Proceeding s of the National Academy of Sciences of the United States of America 1997, 94, (23), 1225812262.
80 39. Sharif, D. I.; Gallon, J.; Smith, C. J.; Dudley, E. Isme Journal 2008, 2, (12), 11711182. 40. An, G. C.; Faeder, J. R. Mathematical Biosciences 2009, 217, (1), 5363. 41. Shabir, S.; Southgate, J. Cell Calcium 2008, 44, (5), 453464. 42. Tanouchi, Y.; Tu, D.; Kim, J.; You, L. Public Library of Science Computational Biology 2008, 4, (8), 8. 43. Goode, B. L.; Drubin, D. G.; Barnes, G. Current Opinion in Cell Biology 2000, 12, (1), 6371. 44. Hess, H. Soft Matter 2006, 2, (8), 669677. 45. Ashkin, A.; Schutze, K.; Dziedzic, J. M.; Euteneuer, U.; Schliwa, M. Nature 1990, 348, (6299), 346348. 46. Greiner, C. M.; Iazikov, D.; Mossberg, T. W. Optics Express 2 006, 14, (25), 1195211957. 47. Pease, R. F.; Chou, S. Y. Proceedings of the I EEE 2008, 96, (2), 248270. 48. Khan, I. A.; Ali, R. Journal of Radiation Research 1985, 26, (1), 109122. 49. Rivera, S. B.; Koch, S. J.; Bauer, J. M.; Edwards, J. M.; Bachand, G. D. Fungal Genetics and Biology 2007, 44, (11), 11701179. 50. Sengupta, S.; Majumder, A. L. Planta 2009, 229, (4), 911929. 51. Cole, M. A.; Voelcker, N. H.; Thissen, H.; Griesser, H. J. Biomaterials 2009, 30, (9), 18271850. 52. Li, N.; Ho, C. M. Lab on a Chip 2008, 8, (12), 21052112. 53. Ionov, L.; Synytska, A.; Diez, S. Advanced Functional Materials 2008, 18, (10), 15011508. 54. Sanii, B.; Parikh, A. N. Annual Review of Physical Chemistry 2008, 59, 411432. 55. Tsai, I. Y.; Crosby, A. J.; Russe ll, T. P., Surface patterning. In Cell Mechanics Elsevier Academic Press Inc: San Diego, 2007; Vol. 83, pp 67 87. 56. Somero, G. N. Annual Review of Ecology and Systematics 1978, 9, (1), 1 29. 57. Somero, G. N. Biochemical Journal 1969, 114, (2), 237&. 5 8. Hazel, J. R.; Prosser, C. L. Physiological Review 1974, 54, (3), 620677. 59. Bouchard, A. M.; Warrender, C. E.; Osbourn, G. C. Physical Review E 2006, 74, (4).
81 60. Hiratsuka, Y.; Miyata, M.; Tada, T.; Uyeda, T. Q. P. Proceedings of the National Academy of Sciences of the United States of America 2006, 103, (37), 13618 13623. 61. Hanson, J.; Huxley, H. E. Nature 1953, 172, (4377), 530532. 62. Svoboda, K.; Schmidt, C. F.; Schnapp, B. J.; Block, S. M. Nature 1993, 365, 721727. 63. Miyamoto, Y.; Muto, E.; Mashimo, T.; Iwane, A. H.; Yoshiya, I.; Yanagida, T. Biophysical Journal 2000, 78, (2), 9409. 64. Tsuda, Y.; Mashimo, T.; Yoshiya, I.; Kaseda, K.; Harada, Y.; Yanagida, T. Biophy sical Journal 1996, 71, (5), 273341. 65. Block, S. M.; Goldstein, L. S.; Sc hnapp, B. J. Nature 1990, 348, (6299), 348 52. 66. Block, S. M. Biophysical Journal 2007, 92, (9), 29862995. 67. Ramachandran, S.; Ernst, K. -H.; Bachand, George D.; Vogel, V.; Hess, H. Small 2006, 2, (3), 330334. 68. RWJU, R. W. J. U. H. abiocor_1.jpg. www.rwjuh.edu/images/news/abiocor_1.jpg (January 28, 2009), 69. Winslow, T.; Kibluk, L. Insulin Production. http://stemcells.nih.gov/info/2006report/2006Chapter7.htm (Jan 28, 2009), 70. Zygote Media Group, I. human heart. http://www.3dscience.com/3D_Models/Human_Anatomy/Heart/index.php (Jan 2 8, 2009), 71. Bequette, B. W. Diabetes Technology and Therapeutics 2005, 7, (1), 2847. 72. Bachand, G. D.; Hess, H.; Ratna, B.; Satir, P.; Vogel, V. Lab on a Chip 2009. 73. Alberts, B., Johnson, A., Lewis, J., Raff, M., Roberts, K. and Walter, P. Molec ular Biology of the Cell. 4th ed.; Garland Science: New York, 2002; Vol. New York. 74. Dechat, T.; Adam, S. A.; Goldman, R. D. Advances in Enzyme Regulation In Press, Corrected Proof. 75. Goldberg, M. W.; Fiserova, J.; Huttenlauch, I.; Stick, R. Biochemical Society Transactions 2008, 36, 13391343. 76. Yoon, M.; Moir, R. D.; Prahlad, V.; Goldman, R. D. Journal of Cell Biology 1998, 143, (1), 147157. 77. Costa, M. L.; Escaleira, R.; Cataldo, A.; Oliveira, F.; Mermelstein, C. S. In Desmin: molecula r interactions and putative functions of the muscle intermediate filament protein 2004; 2004; pp 18191830.
82 78. Bieling, P.; Laan, L.; Schek, H.; Munteanu, E. L.; Sandblad, L.; Dogterom, M.; Brunner, D.; Surrey, T. Nature 2007, 450, (7172), 11001105. 79. Behrens, S.; Rahn, K.; Habicht, W.; Bohm, K. J.; Rosner, H.; Dinjus, E.; Unger, E. Advanced Materials 2002, 14, (22), 16211625. 80. Nogales, E.; Wolf, S. G.; Downing, K. H. Nature 1998, 391, (6663), 199203. 81. Ray, S.; Meyhofer, E.; Milligan, R. A.; Ho ward, J. Journal of Cell Biology 1993, 121, (5), 10831093. 82. Raviv, U.; Nguyen, T.; Ghafouri, R.; Needleman, D. J.; Li, Y.; Miller, H. P.; Wilson, L.; Bruinsma, R. F.; Safinya, C. R. Biophysical Journal 2007, 92, (1), 278287. 83. Mogensen, M. M.; Tucke r, J. B. Journal of Cell Science 1987, 88, 95107. 84. Eichenlaubritter, U. Journal of Cell Science 1985, 76, (JUN), 337355. 85. Pierson, G. B.; Burton, P. R.; Himes, R. H. Journal of Cell Biology 1978, 76, (1), 223228. 86. Don W. Fawcett, K. R. P. Journal of Morphology 1954, 94, (2), 221281. 87. Manton, I.; Clarke, B. Journal of Experimental Botany 1952, 3, (3), 265275. 88. Inoue, S.; Sato, H. Journal of General Physiology 1967, 50, (6), 259292. 89. Inoue, S.; Fuseler, J.; Salmon, E. D.; Ellis, G. W. Biophysical Journal 1975, 15, (7), 725744. 90. Howard, J. Physical Biology 2006, 3, (1), 5466. 91. Hackney, D. D. Proceedings of the National Academy of Sciences of the United States of America 2005, 102, (51), 1833818343. 92. King, S. J.; Schroer, T. A Nature Cell Biology 2000, 2, (1), 2024. 93. Kural, C.; Kim, H.; Syed, S.; Goshima, G.; Gelfand, V. I.; Selvin, P. R. Science 2005, 308, (5727), 14691472. 94. Gross, S. P.; Welte, M. A.; Block, S. M.; Wieschaus, E. F. Journal of Cell Biology 2002, 156, (4), 715724. 95. Schiff, P. B.; Fant, J.; Horwitz, S. B. Nature 1979, 277, (5698), 6657. 96. Snyder, J. P.; Nettles, J. H.; Cornett, B.; Downing, K. H.; Nogales, E. Proceedings of the National Academy of Sciences of the United States of America 2001, 98, (9), 53125316.
83 97. Guo, H. L.; Xu, C. H.; Liu, C. X.; Qu, E.; Yuan, M.; Li, Z. L.; Cheng, B. Y.; Zhang, D. Z. Biophysical Journal 2006, 90, (6), 20932098. 98. Kriss, T. C.; Kriss, V. M. Neurosurgery 1998, 42, (4), 899907. 99. Coons, A. H.; Creech, H. J.; Jones, R. N.; Berliner, E. Journal of Immunology 1942, 45, (3), 159170. 100. Coons, A. H.; Kaplan, M. H. Journal of Experimental Medicine 1950, 91, (1), 1 13. 101. Kaplan, M. H.; Coons, A. H.; Deane, H. W. Journal of Experimental Medicine 1950, 91, (1), 15 30. 102. Chalfie, M.; Tu, Y.; Euskirchen, G.; Ward, W. W.; Prasher, D. C. Science 1994, 263, (5148), 802805. 103. Lakowicz, J. R., Principles of Fluorescence Spectroscopy. 3rd ed.; Springer: Berlin/Heidelberg, 2006; p 954. 104. Howard, J.; Hymann A. A. Nature 2003, 422, 753758. 105. Vale, R. D.; Milligan, R. A. Science 2000, 288, (5463), 8895. 106. Fischer, T.; Agarwal, A.; Hess, H. Nature Nanotechnology 2009, 4, (3), 162166. 107. Khademhosseini, A.; Langer, R.; Borenstein, J.; Vacanti, J. P. Proceedings of the National Academy of Sciences of the United States of America 2006, 103, (8), 24802487. 108. Folch, A.; Toner, M. Annual Review of Biomedical Engineering 2000, 2, 227+. 109. Li, N.; Tourovskaia, A.; Folch, A. Critical Reviews in Biomed ical Engineering 2003, 31, ((5&6)), 423 488. 110. Raghavan, S.; Chen, C. S. Advanced Materials 2004, 16, (15), 13031313. 111. Sniadecki, N.; Desai, R. A.; Ruiz, S. A.; Chen, C. S. Annals of Biomedical Engineering 2006, 34, (1), 5974. 112. Falconnet, D.; Csucs, G.; Grandin, H. M.; Textor, M. Biomaterials 2006, 27, (16), 30443063. 113. Chen, C. S.; Mrksich, M.; Huang, S.; Whitesides, G. M.; Ingber, D. E. Science 1997, 276, (5317), 14251428. 114. McBeath, R.; Pirone, D. M.; Nelson, C. M.; Bhadriraju, K.; C hen, C. S. Developmental Cell 2004, 6, (4), 483495. 115. Senaratne, W.; Andruzzi, L.; Ober, C. K. Biomacromolecules 2005, 6, (5), 24272448.
84 116. Besson, E.; Gue, A. M.; Sudor, J.; Korri Youssoufi, H.; Jaffrezic, N.; Tardy, J. Langmuir 2006, 22, (20), 8346 8352. 117. Babacan, S.; Pivarnik, P.; Letcher, S.; Rand, A. G. Biosensors & Bioelectronics 2000, 15, (11 12), 615621. 118. Feng, C. L.; Embrechts, A.; Bredebusch, I.; Schnekenburger, J.; Domschke, W.; Vancso, G. J.; Schonherr, H. Advanced Materials 2007, 19, (2), 286+. 119. Reuther, C.; Hajdo, L.; Tucker, R.; Kasprzak, A. A.; Diez, S. Nano Letters 2006, 6, (10), 21772183. 120. Allen, R. D. Journal of Photopolymer Science and Technology 2007, 20, (3), 453455. 121. Yamaguchi, M.; Nishimura, O.; Lim, S. H.; Shimokawa, K.; Tamura, T.; Suzuki, M. Colloids and Surfaces a-Physicochemical and Engineering Aspects 2006, 284, 532534. 122. Yamaguchi, M.; Nishimura, O.; Lim, S. H.; Shimokawa, K.; Tamura, T.; Suzuki, M. Colloids and Surfaces A: Physicochemical and Engineering Aspects 2006, 284 285, 532534. 123. Tinazli, A.; Piehler, J.; Beuttler, M.; Guckenberger, R.; Tampe, R. Nature Nanotechnology 2007, 2, (4), 220225. 124. David S. Ginger, H. Z. C. A. M. Angewandte Chemie International Edition 2004, 43, (1) 3045. 125. Xia, Y. N.; Whitesides, G. M. Angewandte Chemie -International Edition 1998, 37, (5), 551575. 126. Slocik, J. M.; Beckel, E. R.; Jiang, H.; Enlow, J. O.; Zabinski, J. S.; Bunning, T. J.; Naik, R. R. Advanced Materials 2006, 18, (16), 2095210 0 127. Jung, J. M.; Kwon, K. Y.; Ha, T. H.; Chung, B. H.; Jung, H. T. Small 2006, 2, (8 9), 10101015. 128. Slocik, J. M.; Beckel, E. R.; Jiang, H.; Enlow, J. O.; Zabinski, J. S.; Bunning, T. J.; Naik, R. R. Advanced Materials 2006, 18, (16), 20952100. 1 29. Holden, M. A.; Cremer, P. S. J ournal of the American Chemical Society 2003, 2003, (125), 80748075. 130. Gerhardt, K. E.; Wilson, M. I.; Greenberg, B. M. Photochemistry and Photobiology 2005, 81, (5), 10618. 131. Ionov, L.; Stamm, M.; Diez, S. Nano Le tters 2006, 6, (9), 19827. 132. Huber, D. L.; Manginell, R. P.; Samara, M. A.; Kim, B.-I.; C. Bunker, B. Science 2003, 301, 352354.
85 133. Indig, G. L.; Jay, D. G.; Grabowski, J. J. Biophysical Journal 1992, 61, (3), 631638. 134. Ionov, L.; Stamm, M.; Die z, S. Nano Lett ers 2006, 6, (9), 19821987. 135. Iyer, K. S.; Klep, V.; Pionteck, J.; Malz, H.; Luzinov, I. Abstracts of Papers of the American Chemical Society 2003, 225, U620-U620. 136. Iyer, K. S.; Zdyrko, B.; Malz, H.; Pionteck, J.; Luzinov, I. Macromolecules 2003, 36, (17), 65196526. 137. Jay, D. G. Proceedings of the National Academy of Sciences of the United States of America 1988, 85, (15), 54545458. 138. Howard, J. Nature 1997, 389, (6651), 561567. 139. Howard, J., Mechanics of Motor Prot eins and the Cytoskeleton. Sinauer Press: Sunderland, Massachusetts, 2001; p 384. 140. Bohm, K. J.; Stracke, R.; Baum, M.; Zieren, M.; Unger, E. FEBS Letters 2000, 466, 5962. 141. Coy, D. L.; Wagenbach, M.; Howard, J. Journal of Biological Chemistry 1999, 274, (6), 36673671. 142. Ionov, L.; Stamm, M.; Diez, S. Nano Lett ers 2005, 5, (10), 19101914. 143. Bezkorovainy, A. R., M.E., Concise Biochemistry New York, 1996. 144. Huber, D. L.; Manginell, R. P.; Samara, M. A.; Kim, B.-I.; Bunker, B. C. Science 200 3, 301, 352354. 145. Guijt, R. M.; Dodge, A.; van Dedem, G. W. K.; de Rooij, N. F.; Verpoorte, E. Lab on a Chip 2003, 3, (1), 1 4. 146. Mihajlovic, G.; Brunet, N. M.; Trbovic, J.; Xiong, P.; von Molnar, S.; Chase, P. B. Applied Physics Letters 2004, 85, ( 6), 10601062. 147. Ruoff, P.; Zakhartsev, M.; Westerhoff, H. V. FEBS Journal 2007, 274, (4), 940950. 148. Horowitz, P., Hill, W., The Art of Electronics Cambridge University Press: Cambridge, 1989. 149. van Schilfgaarde, M.; Abrikosov, I. A.; Johansson, B. Nature 1999, 400, (6739), 4649. 150. Schliwa, M.; Woehlke, G. Nature 2003, 422, (6933), 75965. 151. Howard, J., Mechanics of Motor Proteins and the Cytoskeleton. Sinauer: Sunderland, MA, 2001; p 367.
86 152. Nicolau, D. V.; Suzuki, H.; Mashiko, S.; Tagu chi, T.; Yoshikawa, S. Biophysical Journal 1999, 77, (2), 112634. 153. Hiratsuka, Y.; Tada, T.; Oiwa, K.; Kanayama, T.; Uyeda, T. Q. Biophysical Journal 2001, 81, (3), 155561. 154. Stracke, P.; Bohm, K. J.; Burgold, J.; Schacht, H. J.; Unger, E. Nanotechnology 2000, 11, (2), 52 56. 155. van den Heuvel, M. G. L.; Butcher, C. T.; Lemay, S. G.; Diez, S.; Dekker, C. Nano Letters 2005, 5, (2), 235241. 156. Yokokawa, R.; Takeuchi, S.; Kon, T.; Nishiura, M.; Ohkura, R.; Sutoh, K.; Fujita, H. Journal of Microelectromechanical Systems 2004, 13, (4), 612619. 157. Du, Y. Z.; Hiratsuka, Y.; Taira, S.; Eguchi, M.; Uyeda, T. Q. P.; Yumoto, N.; Kodaka, M. Chemical Communications 2005, 40, (16), 20802082. 158. Huang, Y. M.; Uppalapati, M.; Hancock, W. O.; Jacks on, T. N. I EEE Transactions on Advanced Packaging 2005, 28, (4), 564570. 159. Moorjani, S. G.; Jia, L.; Jackson, T. N.; Hancock, W. O. Nano Letters 2003, 3, (5), 633637. 160. Lin, C. T.; Kao, M. T.; Kurabayashi, K.; Meyhfer, E. Small 2006, 2, (2), 2812 87. 161. Dennis, J. R.; Howard, J.; Vogel, V. Nanotechnology 1999, 10, 232236. 162. Hess, H.; Clemmens, J.; Qin, D.; Howard, J.; Vogel, V. Nano Letters 2001, 1, (5), 235239. 163. Hess, H.; Howard, J.; Vogel, V. Nano Letters 2002, 2, (10), 11131115. 164. Hess, H.; Clemmens, J.; Howard, J.; Vogel, V. Nano Letters 2002, 2, (2), 113116. 165. Dinu, C. Z.; Opitz, J.; Pompe, W.; Howard, J.; Mertig, M.; Diez, S. Small 2006, 2, (8 9), 10901098. 166. Nicolau, D. V.; Nicolau, D. V.; Solana, G.; Hanson, K. L.; Fil ipponi, L.; Wang, L. S.; Lee, A. P. In Molecular motors -based micro and nano-biocomputation devices 2006; 2006; pp 15821588. 167. Bachand, George D.; Rivera, Susan B.; Carroll Portillo, A.; Hess, H.; Bachand, M. Small 2006, 2, (3), 381385. 168. Lin, C. T.; Kao, M. T.; Kurabayashi, K.; Meyhofer, E. Nano Letters 2008, 8, (4), 10411046. 169. Nitta, T.; Hess, H. Nano Letters 2005, 5, (7), 13371342.
87 170. Bhm, K. J.; Stracke, R.; Baum, M.; Zieren, M.; Unger, E. FEBS Letters 2000, 466, (1), 5962. 171. Kawa guchi, K.; Ishiwata, S. Cell Motil Cytoskeleton 2001, 49, (1), 417. 172. Scopes, R. K. Clinica Chimica Acta 1995, 237, (12), 17 23. 173. Kawaguchi, K.; Ishiwata, S. Biochemical and Biophysical Research Communications 2000, 272, (3), 895899. 174. Leduc, C.; Ruhnow, F.; Howard, J.; Diez, S. Proceedings of the National Academy of Sciences of the United States of America 2007, 104, (26), 1084710852. 175. Agarwal, A.; Katira, P.; Hess, H. Nano Letters 2009, 9, (3), 11701175. 176. Bishop, J. D.; Klavi ns, E. Nano Letters 2007, 7, (9), 25742577. 177. Niemeyer, C. M.; Mirkin, C. A., Nanobiotechnology Wiley -VCH: Weinheim, 2004; p 469. 178. Alkon, D. L.; Rasmussen, H. Science 1988, 239, (4843), 9981005. 179. Berridge, M. J.; Bootman, M. D.; Roderick, H. L. Nature Reviews Molecular Cell Biology 2003, 4, (7), 517529. 180. Kindzelskii, A. L.; Petty, H. R. Proceedings of the National Academy of Sciences of the United States of America 2002, 99, (14), 92079212. 181. Alberts, B.; Johnson, A.; Lewis, J.; Raff, M.; Roberts, K.; Walter, P., Molecular Biology of the Cell. Fourth Edition. 4 ed.; Garland: New York, 2002. 182. Balzani, V. V.; Credi, A.; Raymo, F. M.; Stoddart, J. F. Angewandte Chemie International Edition 2000, 39, (19), 33483391. 183. Yan, H.; Zhang, X.; Shen, Z.; Seeman, N. C. Nature 2002, 415, (6867), 625. 184. Namasivayam, V.; Larson, R. G.; Burke, D. T.; Burns, M. A. Analytical Chemistry 2003, 75, (16), 41884194. 185. Hess, H.; Bachand, G. D.; Vogel, V. Chemistry A European Journal 2004, 10, (9), 21102116. 186. Hess, H.; Vogel, V. Reviews in Molecular Biotechnology 2001, 82, (1), 6785. 187. de Graaf, R. A.; van Kranenburg, A.; Nicolay, K. Biophysical Journal 2000, 78, (4), 16571664. 188. Howard, J.; Hudspeth, A. J.; Vale, R. D. Nature 1989, 342, 154158.
88 189. Schief, W. R.; Clark, R. H.; Crevenna, A. H.; Howard, J. Proceedings of the National Academy of Sciences of the United States of America 2004, 101, (5), 11831188. 190. Klumpp, S.; Lipowsky, R. Proceedings of the National Academy of Sc iences of the United States of America 2005, 102, (48), 1728417289. 191. Bronshtein, I. N.; Semendyayev, K. A.; Musiol, G.; Muehlig, H., Handbook of mathematics 4th ed.; Springer: Berlin, 2004. 192. Wu, D.; Tucker, R.; Hess, H. IEEE Transactions on Advanced Packaging 2005, 28, (4), 594599. 193. Beta, C.; Wyatt, D.; Rappel, W. J.; Bodenschatz, E. Anal ytical Chemistry 2007, 79, (10), 39403944. 194. Liu, H.; Schmidt, J. J.; Bachand, G. D.; Rizk, S. S.; Looger, L. L.; Hellinga, H. W.; Montemagno, C. D. Nature Materials 2002, 1, (3), 173177. 195. Yurke, B.; Turberfield, A. J.; Mills, A. P.; Simmel, F. C.; Neumann, J. L. Nature 2000, 406, (6796), 605608. 196. Bath, J.; Turberfield, A. J. Nature Nanotechnology 2007, 2, (5), 275284. 197. Hirabayashi, M.; Taira, S.; Kobayashi, S.; Konishi, K.; Katoh, K.; Hiratsuka, Y.; Kodaka, M.; Uyeda, T. Q. P.; Yumoto, N.; Kubo, T. Biotechnology and Bioengineering 2006, 94, (3), 473480. 198. Taira, S.; Du, Y. Z.; Hiratsuka, Y.; Konishi, K.; Kubo, T .; Uyeda, T. Q. P.; Yumoto, N.; Kodaka, M. Biotechnology and Bioengineering 2006, 95, (3), 533538. 199. Huber, D. L.; Manginell, R. P.; Samara, M. A.; Kim, B. I.; Bunker, B. C. Science 2003, 301, (5631), 3524. 200. Kocer, A.; Walko, M.; Meijberg, W.; Fer inga, B. L. Science 2005, 309, (5735), 755758. 201. Zhu, Y.; Fujiwara, M. Angewandte Chemie International Edition 2007, 46, (13), 22412244. 202. Martin, B. D.; Velea, L. M.; Soto, C. M.; Whitaker, C. M.; Gaber, B. P.; Ratna, B. Nanotechnology 2007, 18, ( 5). 203. Riveline, D.; Ott, A.; Julicher, F.; Winkelmann, D. A.; Cardoso, O.; Lacapere, J. J.; Magnusdottir, S.; Viovy, J. L.; Gorre Talini, L.; Prost, J. European Biophysics Journal 1998, 27, (4), 403408. 204. Bhm, K. J.; Stracke, R.; Mhlig, P.; Unger, E. Nanotechnology 2001, 12, 238244. 205. van den Heuvel, M. G. L.; De Graaff, M. P.; Dekker, C. Science 2006, 312, (5775), 910914.
89 206. Kim, T.; Kao, M. T.; Hasselbrink, E. F.; Meyhofer, E. Nano Letters 2007, 7, (1), 211217. 207. Kim, T.; Kao, M. T.; M eyhofer, E.; Hasselbrink, E. F. Nanotechnology 2007, 18, (2). 208. Hutchins, B. M.; Platt, M.; Hancock, W. O.; Williams, M. E. Small 2007, 3, (1), 126131. 209. Ionov, L.; Stamm, M.; Diez, S. Nano Letters 2006, 6, (9), 19821987. 210. Hofstadter, D. R., G del, Escher, Bach Vintage Books Edition ed.; Random House: New York, 1980; p 777. 211. Wikswo, J. P.; Prokop, A.; Baudenbacher, F.; Cliffel, D.; Csukas, B.; Velkovsky, M. Iee Proceedings -Nanobiotechnology 2006, 153, (4), 81101. 212. Nakano, T.; Suda, T. Applied Soft Computing 2007, 7, (3), 870878. 213. Helbing, D.; Deutsch, A.; Zerial, M.; Schulze, F.; Diez, S.; Breier, G.; Peters, K., Ladenburger Discourse: From BioInspired Logistics to Logistics Inspired Bio Nano Engineering. In Hess, H., Ed. 2007. 214. Clemmens, J.; Hess, H.; Lipscomb, R.; Hanein, Y.; Boehringer, K. F.; Matzke, C. M.; Bachand, G. D.; Bunker, B. C.; Vogel, V. Langmuir 2003, 19, (26), 1096710974. 215. Montemagno, C.; Bachand, G. Nanotechnology 1999, 10, 225331. 216. Agui, L.; Eguilaz, M.; Pena -Farfal, C.; Yanez -Sedeno, P.; Pingarron, J. M. In Lactate Dehydrogenase Biosensor Based on an Hybrid Carbon Nanotube -Conducting Polymer Modified Electrode 2009; Wiley-V C H Verlag Gmbh: 2009; pp 386391. 217. York, J.; Spetzler, D.; Xiong, F. S.; Frasch, W. D. Lab on a Chip 2008, 8, (3), 415419. 218. Block, S. M. Cell 1996, 87, (2), 151157. 219. Schnitzer, M. J.; Block, S. M. Nature 1997, 388, (6640), 38690. 220. Spudich, J. A.; Kron, S. J.; Sheetz, M. P. Nature 1985, 315, 584586. 221. Toyoshima, Y. Y.; Kron, S. J.; Spudich, J. A. Proceedings of the National Academy of Sciences of the United States of America 1990, 87, (18), 71307134. 222. Hodge, T.; Cope, M. J. Journal of Cell Science 2000, 113, (19), 33533354. 223. Yildiz, A.; Forkey, J. N.; McKinney, S. A.; Ha, T.; Goldman, Y. E.; Selvin, P. R. Science 2003, 300, (5628), 20615. 224. Iwaki, M.; Iwane, A. H.; Ikebe, M.; Yanagida, T. In Biased Brownian motion mechanism for processivity and directionality of single -headed myosin-VI 2008; Elsevier Sci Ltd: 2008; pp 3947.
90 225. Rios, L.; Bachand, G. D. Lab on a Chip 2009, 9, (7), 10051010. 226. Buehler, M. J.; Yung, Y. C. Nature Materials 2009, 8, (3), 175188. 227. Hunt, A. J.; Howard, J. Proceedings of t he National Academy of Sciences of the United States of America 1993, 90, (24), 116537. 228. Janson, M. E.; Dogterom, M. Biophysical Journal 2004, 87, (4), 27232736. 229. Pampaloni, F.; Lattanzi, G.; Jon, A.; Surrey, T.; Frey, E.; Florin, E. -L. Proce edings of the National Academy of Sciences of the United States of America 2006, 103, (27), 1024810253. 230. Van den Heuvel, M. G. L.; de Graaff, M. P.; Dekker, C. Proceedings of the National Academy of Sciences of the United States of America 2008, 105, (23), 79417946. 231. Boal, Andrew K.; Tellez, H.; Rivera, Susan B.; Miller, Nicholas E.; Bachand, George D.; Bunker, Bruce C. Small 2006, 2, (6), 793803. 232. Turner, D.; Chang, C.; Fang, K.; Cuomo, P.; Murphy, D. Analytical Biochemistry 1996, 242, (1), 2025. 233. Doot, R. K.; Hess, H.; Vogel, V. Soft Matter 2007, 3, (3), 349356. 234. Bohm, K. J.; Beeg, J.; zu Horste, G. M.; Stracke, R.; Unger, E. I EEE Transactions on Advanced Packaging 2005, 28, (4), 571576. 235. Limberis, L.; Stewart, R. J. Nanotechnology 2000, 11, (2), 4751. 236. Yokokawa, R.; Yoshida, Y.; Takeuchi, S.; Kon, T.; Fujita, H. Nanotechnology 2006, 17, 289. 237. Yokokawa, R.; Murakami, T.; Sugie, T.; Kon, T. Nanotechnology 2008, 19, (12), 125505. 238. Yokokawa, R.; Yoshida, Y.; Takeuchi, S.; Kon, T.; Sutoh, K.; Fujita, H. IEEE Transactions on Advanced Packaging 2005, 28, (4), 577583. 239. Tucker, R.; Ionov, L.; Diez, S. manuscript in preparation 2009. 240. Tucker, R.; Saha, A. K.; Katira, P.; Bachand, M.; Bachand, G. D.; Hess, H Small 2009, X, (X), 1 4. 241. Tucker, R.; Katira, P.; Hess, H. Nano Letters 2008, 8, (1), 221 226. 242. Tucker, R.; Hess, H. manuscript in preparation 2009. 243. Schoch, R. B.; Han, J.; Renaud, P. Reviews of Modern Physics 2008, 80, (3), 839.
91 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 WashingtonSeattle. Rob c ontinued 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, dev elop, and produce biomedical devices and point of care diagnostic systems using hybrid bionanotechnologies.