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Investigating nanomechanical properties of proteins that exhibit differing functionality using single… Jollymore, Ashlee 2009

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INVESTIGATING NANOMECHANICAL PROPERTIES OF PROTEINS THAT EXHIBIT DIFFERING FUNCTIONALITY USING SINGLE MOLECULE ATOMIC FORCE MICROSCOPY  by  ASHLEE JOLLYMORE BSc., McGill University, 2006  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE  in  THE FACULTY OF GRADUATE STUDIES (Chemistry)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2009  ©Ashlee Jollymore, 2009  Abstract Proteins are the workhorses of the biological world, performing a range of tasks that are critical to life. How proteins attain their functional three dimensional fold from a simple linear polymer chain is a poorly understood, yet fundamental process. Comprehending how proteins fold and the molecular arbiters of functional characteristics has the potential for the future rational engineering of proteins that exhibit desired utility and remains an active area of research today. The atomic force microscope (AFM) has been developed as a tool uniquely suited towards studying protein folding and functionality at the nanoscale, with its ability to definitively describe the mechanical properties of a single molecule of protein. This study seeks to apply the AFM towards elucidating the mechanical properties of two proteins with dissimilar functionality: the mechanical extracellular matrix protein tenascin-X and the non-mechanical enzyme bacillus circulans xylanase (BCX). By explicating the mechanical properties of each of these proteins, we seek to explore how intrinsic functionality is encoded within the molecular structure of a protein with intrinsic mechanical functionality as well as one lacking a mechanical physiological role. Using AFM to investigate these differences expands on what is already known concerning the mechanical properties of proteins exhibiting differing structure and functional character, seeking to further explicate the basis of mechanical properties observable using the AFM.  ii  TABLE OF CO TE TS Abstract ........................................................................................................................................... ii Table of Contents ........................................................................................................................... iii List of Tables ................................................................................................................................ vii List of Figures .............................................................................................................................. viii Acknowledgements ........................................................................................................................ ix Dedication ....................................................................................................................................... x Co-Authorship Statement............................................................................................................... xi Chapter 1 Introduction .................................................................................................................... 1 1.1 Elucidating the Protein Folding Mechanism is a Viable and Unsolved Biological Question .................................................................................................................................... 1 1.1.1 Introduction to Proteins: How Folding Determines Critical Functionality ............... 1 1.1.2 The Protein Fold: Primary, Secondary, Tertiary and Quaternary Structure .............. 1 1.1.3 Protein Folding is a Directed Process Driven by a Number of Forces ...................... 2 1.1.3.1 Conformational Free Energy and the Rough Energy Landscape................ 3 1.1.4 Protein Folding is Encoded Within the Key Residues in the Primary Sequence....... 4 1.1.5 How Molecular Characteristics Determine Mechanical Properties: Mechanical vs. Non-Mechanical Proteins .............................................................................................. 6 1.2 Understanding Protein Folding ........................................................................................... 8 1.2.1 Methods of Elucidating Protein Folding .................................................................... 8 1.2.1.1 Methods of Determining Folded Structure ................................................. 9 1.2.1.2 Ensemble Methods of Elucidating Protein Folding and Unfolding .......... 10 1.2.1.3 Single Molecule Methods of Studying Protein Folding and Unfolding: FRET ..................................................................................................................... 11 1.2.2 Single Molecule Force Spectroscopies .................................................................... 12 1.2.2.1 Force as a Physiologically Relevant Means of Unfolding ........................ 12 1.2.2.2 Force Spectroscopy Techniques ............................................................... 13 1.2.2.2.1 Optical Tweezers ................................................................................ 13 1.2.2.2.1 Magnetic Tweezers ............................................................................. 15 1.2.2.3 The Atomic Force Microscope .............................................................. 15 1.2.2.3.1 Origins and Technological Advancements of the Atomic Force Microscope ......................................................................................................... 15 iii  1.3 The Atomic Force Microscope ......................................................................................... 19 1.3.1 Overview of the Atomic Force Microscope............................................................. 19 1.3.2 The Piezoelectric Positioner and Glass Coverslip ................................................... 20 1.3.3 The AFM Head ........................................................................................................ 22 1.3.4 Cantilever Calibration and the Equipartition Theorem ............................................ 24 1.3.5 The Optical Detection System ................................................................................. 25 1.3.6 Piezoelectric Positioner Driver and Output Software .............................................. 25 1.3.7 Experimentally Pulling a Single Protein Molecule .................................................. 26 1.3.7.1 Force Clamp Experiments......................................................................... 26 1.3.7.1 Constant Speed Experiments .................................................................... 26 1.4 Investigating the Nanomechanical Fingerprint of Mechanical Unfolding ....................... 27 1.4.1 General Features of a Force Extension Curve ......................................................... 27 1.4.2 Improving Features of the Force Extension Curve Using Polyprotein Constructs .. 30 1.4.3 Entropic Restoring Forces and the Worm Like Chain Model ................................. 31 1.4.3.1 Persistence Length as a Measure of Polymer Stiffness ............................ 31 1.4.3.2 Contour Length and the Experimental Contour Length Increment .......... 32 1.4.4 Mechanical Unfolding and Unfolding Kinetics of Proteins .................................... 33 1.4.4.1 Exploring the Kinetics of the Folded to Unfolded Transition Under Force ..................................................................................................................... 34 1.4.4.2 The Non-Equilibrium Nature of Unfolding Under Force ......................... 38 1.4.4.3 Non-Equilibrium Implications for Free Energy Calculation .................... 38 1.4.4.4 The Dependence of Unfolding Force on Pull Speed and the Calculation of Unfolding Kinetics ........................................................................ 39 1.4.4.5 Fitting Unfolding Force Distribution Using Unfolding Kinetic Parameters ............................................................................................................. 40 1.4.4.6 Mechanical Stability is a Kinetic Rather Than Thermodynamic Parameter .............................................................................................................. 41 1.5 The Molecular Determinates of Mechanical Stability as Determined using Atomic Force Spectroscopy ................................................................................................................. 41 1.5.1 Emerging Trends in Structural Determinates of Mechanical Behaviour ................. 41 1.5.2 Investigating The Nanomechanical Properties of a Mechanical and NonMechanical Proteins using Single Molecule AFM ........................................................... 45 1.6 References ......................................................................................................................... 47 Chapter 2 The Nanomechanical Properties of Tenascin-X........................................................... 53 2.1 Introduction to the Tenascins: A Family of Ubiquitous Extracellular Membrane Proteins ................................................................................................................................... 53 2.1.1 The Extracellular Matrix: A Complex Matrix with Probable Force Transduction Functionality ..................................................................................................................... 53 2.1.2 The Tenascin Family and Their Importance in ECM Organization Through Cell Adhesion ........................................................................................................................... 54 2.1.3 Tenascin-X is An Integral Connective Protein Within the ECM ............................ 55 2.1.4 Tenascin-X Structure ............................................................................................... 55 iv  2.1.5 Tenascin-X as a Force Spectroscopy Model System ............................................... 56 2.2 The Nanomechanical Properties of Tenascin-X as Demonstrated using Single Molecule AFM ........................................................................................................................ 57 2.2.1 The Unfolding Behaviour of Full Length TN-X ..................................................... 57 2.2.1.1 Pinpointing Molecular Origins of Mechanical Features: rTNX∆E∆F ...... 58 2.2.1.2 Determining the Unfolding Force and Contour Length Increment of rTNX∆E∆F ........................................................................................................... 59 2.2.1.3 Speed Dependence of Unfolding Forces Within FnIII Domains .............. 62 2.2.1.4 Folding Kinetics of FnIII Domains ........................................................... 63 2.2.2 Mechanical Unfolding and Folding Dynamics of Individual FnIII Domains of Tenascin-X ........................................................................................................................ 65 2.2.2.1 The Mechanical Unfolding Properties of TNXfn10 ................................. 68 2.2.2.2 The Mechanical Properties of TNXfn11................................................... 69 2.3 Discussing Tenascin-X as an Elastic Protein with Well Defined Mechanical Properties ................................................................................................................................ 71 2.3.1 Comparison of the Mechanical Features of FnIII Domains Between Tenascin-X and Tenascin-C ................................................................................................................. 74 2.4 Conclusion ........................................................................................................................ 76 2.5 Investigating the Properties of TN-X: Materials and Methods ......................................... 77 2.5.1 Full Length and All FnIII TN-X Fragment .............................................................. 77 2.5.2 Construction of FnIII Containing Polyproteins ....................................................... 77 2.5.2.1 Construction of Genes Containing FnIII10 and FnIII11........................... 77 2.5.2.2 Expression of the FnIII Containing Polyprotein ....................................... 80 2.5.3 Single Molecule AFM Experiments ........................................................................ 80 2.6 References ......................................................................................................................... 82 Chapter 3 Investigation of the Nanomechanical Properties of a Non-Mechanical Protein: Bacillus Circulans Xylanase ......................................................................................................... 85 3.1 Introduction to Bacillus Circulans Xylanase .................................................................... 85 3.1.1 BCX is a Bacterial Enzyme Responsible for the Breakdown of Cell Walls ........... 85 3.1.2 Variability within the Structure of Xylan Has Resulted in a Multiplicity of Xylanase Forms ................................................................................................................ 86 3.1.3 Investigating Xylanases as a ‘Greener’ Bleaching Agent Within Paper Industries. 87 3.1.4 BCX Structure .......................................................................................................... 88 3.1.5 BCX Mechanism: How Xylan Cleavage is Accomplished ..................................... 89 3.2 BCX as a Force Spectroscopy Model System .................................................................. 91 3.2.1 Previous Studies on the Thermal Stability of BCX ................................................. 92 3.3 Investigating the Mechanical Properties of BCX ............................................................. 95 v  3.3.1 Research Overview .................................................................................................. 95 3.3.2 Investigating the Mechanical Properties of BCX using Single Molecule AFM ...... 95 3.3.2.1 The Pull Rate Dependence of BCX Determines its Unfolding Rate Parameters ........................................................................................................... 101 3.3.2.2 Refolding of BCX Reveals Folding Kinetics.......................................... 102 3.3.2.3 The Unfolding of Wild Type BCX Reveals its Nanomechanical Behaviour ............................................................................................................ 103 3.3.3 Determining the Mechanical Effects of Inhibitor Binding .................................... 106 3.3.3.1 Examining the Effects of Inhibitor Binding on Unfolding Force ........... 108 3.3.4 Investigating the Effects of Structural Changes on the Mechanical Behaviour of BCX ................................................................................................................................ 109 3.3.4.1 The Mechanical Effects of Introduced Bonding: Engineered Disulfide Bonds .................................................................................................................. 111 3.4 Conclusion ...................................................................................................................... 112 3.5 Materials and Methods .................................................................................................... 113 3.5.1 Construction of BCX-Containing Proteins ............................................................ 113 3.5.1.1 Construction of the (GB1)4BCX(GB1)4 Plasmid ................................... 113 3.5.1.1.1 Protein Expression of (GB1)4BCX(GB1)4........................................... 114 3.5.1.2 Construction of Disulfide Bonded Mutants ............................................ 115 3.5.2 AFM Experiments .................................................................................................. 116 3.5.3 Inhibitor Binding to BCX for AFM Experiments .................................................. 116 3.6 References ....................................................................................................................... 118 Chapter 4 Comparing the Naomechanical Properties of Mechanical and Non-Mechanical Proteins: Elucidating the Molecular Determinants of Mechanical Attributes ............................ 122 4.1 Investigating Non-Mechanical and Mechanical Proteins Using Single Molecule AFM as a Means of Illuminating Origins of Mechanical Behavior ............................................... 122 4.2 Future Studies of Protein Nanomechanics ...................................................................... 125 4.3 References ....................................................................................................................... 127  vi  LIST OF TABLES Table 1.1 Mechanical Properties Previously Elucidated using Single Molecule AFM..............................45 Table 3.1 Primers Used in the Construction of Disulfide Mutants............................................................116  vii  LIST OF FIGURES Figure 1.1 Figure 1.2 Figure 1.3 Figure 1.4 Figure 1.5 Figure 1.6 Figure 1.7 Figure 1.8  Rough Energy Landscape Inscribed by Protein Folding ....................................................4 Mechanical Adaptations of the Muscle Protein Titin .........................................................8 Schematic of the AFM Instrument ...................................................................................19 Unfolding a Modular Protein Results in a Representative Sawtooth Pattern ....................29 Representations of the Contour Length and ∆Lc ...............................................................33 The Free Energy Representation of Two State Folding/Unfolding ...................................35 Energetic Changes Caused by Application of Force ........................................................36 Structural Trends in Mechanical Stability .........................................................................42  Figure 2.1 Figure 2.2 Figure 2.3 Figure 2.4 Figure 2.5 Figure 2.6 Figure 2.7 Figure 2.8 Figure 2.9 Figure 2.10 Figure 2.11 Figure 2.12 Figure 2.13 Figure 2.14  Bovine Tenascin-X Structure ............................................................................................56 The Force Extension Fingerprint of Full Length Tenascin-X ...........................................58 The Force Extension Characteristics of rTNX∆E∆F.........................................................59 The Unfolding Force Histogram of Full Length Tenascin-X ............................................60 The Contour Length Increment Histogram of Full Length Tenascin-X ............................61 Mechanical Properties of rTNX∆E∆F ...............................................................................62 Unfolding Kinetics of Full Length and rTNX∆E∆F .........................................................63 Folding Kinetics of Full Length Tenascin-X.....................................................................65 Mechanical Unfolding of FnIII10 and FnIII11 Polyproteins ............................................67 Unfolding Force Histogram and Kinetics of TNXfn10 .....................................................68 Folding Kinetics of TNXfn10 ...........................................................................................69 The Unfolding Force and Unfolding Kinetics of FNXfn11 ..............................................70 Folding Kinetics of TNXfn11 ...........................................................................................71 The Folding Fidelity of Tenascin-X is Greater than Tenascin-C ......................................76  Figure 3.1 Figure 3.2 Figure 3.3 Figure 3.4 Figure 3.5 Figure 3.6 Figure 3.7 Figure 3.8 Figure 3.9 Figure 3.10 Figure 3.11  The Xylan Backbone: The Substrate for BCX ..................................................................87 The Structure of Bacillus Circulans Xylanase (BCX) ......................................................89 BCX Cleaves Xylans Through A Double Displacement Mechanism ...............................91 Inhibitor Binding Forms a Long-Lived Covalent Species.................................................94 The BCX Polyprotein and its Resultant Force Extension Fingerprint ..............................97 Force Histogram and Contour Length Increment of BCX ................................................99 Unfolding Force Dependence on Pulling Speed: Unfolding Kinetics .............................101 The Folding Kinetics of BCX .........................................................................................103 Unfolding of Inhibitor Bound BCX ................................................................................108 Investigating the Force Extension Fingerprint of BCX100x148 .....................................110 The Average Unfolding Force and ∆Lc of BCX100x148 ...............................................111  viii  Acknowledgements The author would like to firstly thank Dr. Hongbin Li for his patience and guidance over the course of this work. The passion and ambition that he brings to his work is perceptible, and I greatly appreciate the effort, creativity and energy that he consistently shares with those around him. I have learned an immeasurable amount during my time in his lab, and am grateful for the opportunities that Dr. Li has given. I would also like to thank the past and present members of the Li lab for their patience and willingness to answer any question or help with any problem encountered on a day to day basis. Although all members of the Li lab have been more than willing to give moral or technical support during my time here, I would especially like to single out Yi Cao, Qing Peng and Deepak Sharma for their infinite patience in teaching me the technical skills necessary for this work as well as their continued support. Despite their own busy schedules, they were always more than willing to sacrifice their own time to aid me and others within the lab. I wish them only the best in their future endeavours, and hope that they achieve the success they rightly deserve. The author would also like to thank the various collaborators and members of the UBC community that made the research presented here possible. Dr. Claire Lethias graciously provided tenascin-X primers used in Chapter Two, while Dr. Lawrence McIntosh provided primers containing the BCX gene used in Chapter Three. Dr. Stephen G. Withers provided the BCX inhibitor used, and Dr. Thomas Morley from Dr. Withers’s lab also provided much appreciated information regarding the proper use of this inhibitor. The author would also like to thank those associated with the Biological Services Facility for their help and for maintaining a facility critical to the research presented here. Lastly, I would like to thank my parents for always being there.  ix  Dedication For my parents, Buzz, my G&G, and my P.  x  Co-Authorship Statement Portions of this thesis benefited from the intellectual and experimental contribution of others. In chapter two, Hongbin Li was responsible for the experimental design of the project as well as the writing of the published manuscript. Yi Cao and Qing Peng both contributed experimentally, provided guidance for the author’s experimental work and helped in editing the published manuscript. Clairie Lethias graciously provided the original tenascin-X plasmids, provided input to helped shape the experimental design of the project, and aided in editing the published manuscript. The author provided the bulk of the experimental work as well as contributions to the written published manuscript. Chapter two is based on the published manuscript, but was written by the author. In chapter three, Hongbin Li was again responsible for the experimental design of the project. The original BCX plasmid was generously provided by Lawrence McIntosh and the inhibitor graciously given by Steven Withers. The author was responsible for carrying out experiments as well as contributions to the experimental design of the project.  xi  Chapter 1 Introduction 1.1 Elucidating the Protein Folding Mechanism is a Viable and Unsolved Biological Question 1.1.1 Introduction to Proteins: How Folding Determines Critical Functionality Proteins are critical to biological life: they act as structural elements, forming intricate structures that form tissues such as muscles, they generate dynamic forces, as in the case of myosin and kinesin, and perform a host of activities upon binding substrate molecules. Proteins are the basis for a host of cellular processes vital to biological functionality, such as energy metabolism, nerve conduction, hormone synthesis and regulation, and even disease. This incredible multiplicity of protein functionality stems from structural diversity, whereby this structure is intrinsic to the protein’s biological functionality. This diversity is huge, from simple arrangements of ß-sheets and α-helices to finely designed jelly-roll topologies to huge multidomain assemblies. That this remarkable assortment stems from arrangements of simple linear polymers delineates the subtlety and sophistication of natural nanomechanical design. How proteins attain their specific, functional fold from the primary amino acid sequence is a poorly understood process, yet understanding the forces that act to drive proteins into their observed functional fold is critical to understanding protein mechanics and dynamics. Potential applications could include engineering proteins with desired functionality as well as further understanding the many diseases that stem from protein misfolding. 1.1.2 The Protein Fold: Primary, Secondary, Tertiary and Quaternary Structure All proteins begin with a primary sequence composed of 20 amino acids bonded through a nitrogen-carbon peptide bond. This primary structure is arranged into secondary structure composed of ß-sheets and α-helices, which are further arranged into tertiary structure from complex combinations of secondary structure. These can in turn form complex multidomain quaternary structures to give the final overall architecture of a protein. Specificity for type is found at each level of structural organization and is, in a sense, cumulative. A certain type of protein, such as the protein form GB1, will exhibit the same 1  primary structure, which in turn leads to folding into similar higher order structure. Significant alteration at this primary level affects the entire organization of structure, where the shape that one level achieves depends on the last and so forth. The final culmination of all of these levels of structural organization results in the final folded form of the protein, and it is this conclusive shape that dictates the biological utility of the particular protein type. The correct functionality is entirely dependent on the precision of this fold; a multitude of disease such as certain cancers, Alzheimer’s and Creutzfeldt-Jakob disease are all due to some form of protein misfolding. That this critical final structure is encoded within every level of organization, beginning from the simply arrayed primary structure, is also easily apparent. However, simply stating that the primary sequence is the arbiter of protein folding, which in turn determines how the protein functions, is a drastic and misleading oversimplification of a difficult question. 1.1.3 Protein Folding is a Directed Process Driven by a umber of Forces From a physical standpoint, proteins fold due to a balance of forces that narrowly favor the folded state over the unfolded polypeptide chain. This preferential native state lies at a thermodynamically stable position relative to that of the unfolded state, which is separated from the native state by a large activation energy barrier. Because of the stable position of the folded state and the size of the activation barrier, proteins tend to fold spontaneously and stay folded under ambient conditions. This process of folding is a directed process; if folding occurred simply by chance, the time taken by each residue to sample its conformational space would be enormous. This impossibility was identified over 50 years ago, a time in which protein studies were still in their infancy, and forms the basis of the Levinthal Paradox. The biological utility of most proteins depends on their ability to fold correctly and quickly; thus, protein folding is a driven process that occurs on a timescale ranging from microseconds to seconds. A number of forces are involved in driving the folding process. Most significant are hydrophobic forces that derive from the presence of hydrophobic amino acids within the protein chain, whereby protein folding occurs to protect these residues from solvent exposure by formation of a hydrophobic core. The protection of hydrophobic side chains from solvent results in the ‘hydrophobic collapse’ of the partially folded protein into a molten globular structure observed by circular dichroism spectroscopy 2  in the folding of certain proteins under particular conditions.1-3 Also contributing to folded stability is hydrogen bonding and ion pairing between amino acids and intrinsic properties of the primary sequence, which helps to confer specific secondary structure. Often, the thermodynamic stability of a folded structure is only marginally greater than that of an unfolded protein, as folding is actually opposed in a number of ways. The largest contributing destabilizing force is the decrease in entropy experienced by individual amino acids that occurs upon folding of a peptide chain into a constrained, compact state. Although other forces, such as the increase in electrostatic charge density that occurs upon folding, act to destabilize the native fold, it is this loss of entropy upon folding that is the main force acting against folding. 1.1.3.1 Conformational Free Energy and the Rough Energy Landscape The correctly folded structure of a protein represents a tiny percentage of the total number of conformations possible based on the complexity and size of most proteins. This small number of ensembles represents stable conformations that result in the greatest decrease in free energy when compared to unfolded, misfolded and intermediate conformations. Thus, native conformations are those that result in the greatest reduction of free energy. If all possible conformations are compared on the basis of their relative energy, a rough energy landscape results.4-7 The ‘roughness’ of this landscape corresponds to the presence of a number of local minima, where the native state is the narrow ensemble of structures that lie at a global minimum. In terms of this energy landscape, unfolded proteins begin at some high-energy conformation. As folding progresses, the protein negotiates its energy landscape, inscribing a downward path of intermediate states, kinetic traps and multiple conformations enroute to the native fold corresponding to the bottom of the energy well. Therefore, protein folding, and conversely unfolding, is an anisotropic process where folding or unfolding may describe a number of paths through the energy landscape. Protein folding and unfolding is highly anisotropic; for example, thermal denaturation of protein from a folded state to unfolded state may go through a different unfolding path than that of chemical or mechanical denaturation.8 In any case, the actual energy difference between unfolded and folded states is usually quite small, and folded proteins are only slightly more thermodynamically stable than their denatured forms. The size of the activation barrier 3  separating the native and denatured states ensures that thermal energy available in the environment is not enough to unfold the majority of molecules within an ensemble. Therefore, unfolding is a probabilistic event that is improbable under normal circumstances, with proteins tending to retain their functional native fold. Figure 1.1 Rough Energy Landscape Inscribed by Protein Folding  Energy Entropy  Figure 1.1 The energy landscape detailing the total conformational space available to a protein and its corresponding energy. Conformations stemming from an unfolded protein exist at high energy, where folding occurs into the narrow range of conformations existing at the folded ‘N’ minima. The rough energy landscape details the kinetic traps, intermediate states, and multiple energy barriers that may occur within the folding mechanism of a protein. Reprinted by permission from Macmillan Publishers Ltd: Nature Structural Biology, Ref(5), copyright 1997.  Kinetically, protein folding is usually represented as a two-state process. Within a two-state process, the folding progresses from denatured states through an energy barrier to the native state.9 Within this model, different unfolded and partially unfolded conformations are in rapid equilibrium with one another and are considered to be one ‘unfolded’ state.10 The predominant model used to describe protein folding is the two-state model, though this may be refined as further kinetic information regarding the protein folding mechanism is elucidated. 1.1.4 Protein Folding is Encoded Within the Key Residues in the Primary Sequence Elucidating the kinetics and thermodynamics behind protein folding is simpler than attempting to predict the final architecture of a protein from its primary sequence. 4  Information concerning the unique structure attained is encoded within the distinctive primary sequence of a protein.11-13 Exactly how structural information is fixed within the primary sequence is confused by the fact that two proteins may exhibit high sequence homology, yet fold into two completely different structures. The alternative is true as well, where domains that originate from two different parent proteins attain similar final folds yet present with low sequence homology, as in the case of fibronectin type III domains.14 This makes the prediction of structure based solely upon sequence homology through methods such as comparative modeling, fold recognition, or de novo prediction methods inexact for some forms of proteins.15 It appears as though the task of folding is shared unequally among residues in a chain, with important amino acids located at key locations along the protein chain. These residues may form essential disulfide or hydrogen bonds,14 or may be larger sequences likely to produce secondary structures such as α-helices or β-sheets. These critical amino acids most likely constitute a relatively small proportion of the entire sequence, participating in folding in a manner difficult to predict. Thus, identification of these key residues is difficult. Identifying the molecular determinates of folded structure would be a major feat in the study of protein mechanics for a number of reasons. Identifying determinates for certain types of structure would impart the ability to design a protein to fold specifically into a form with desired functionality. This is the cornerstone of protein engineering as well as a means of demonstrating proof of concept concerning knowledge about the protein folding mechanism. Although there has been some success in designing proteins with a novel practical fold,16 the protein folding mechanism is not understood well enough to apply this knowledge to the predictable and reliable engineering of functional proteins. Working to further understand the mechanisms behind protein folding has far ranging biological implications. This includes insight into the origins of disease-causing misfolding, to understanding how biological tissues are naturally engineered for specific utility. The delicately arranged titin molecules in muscle, molecular kinesin motors capable of carrying small molecules, and heteropolyproteins made up of individually folded domains meant to unfold when exposed to force demonstrate useful models for attempting to engineer proteins with similar or enhanced functions.  5  1.1.5 How Molecular Characteristics Determine Mechanical Properties: Mechanical vs. on-Mechanical Proteins The manner in which proteins fold determines their structure and in turn establishes their function; thus, ascertaining the molecular basis of how proteins behave mechanistically is intrinsic to the study of how proteins fold. Proteins may be loosely grouped based on the biological functionality they exhibit. A large number of proteins are obviously mechanical in nature; they may be engineered to withstand the force they are subject to physiologically such as titin from muscle sacromeric fibres, be made to transduce force into a biological function, or may also be the originator of force in the manner of molecular motors. Other proteins are apparently non-mechanical in their function, acting to catalyze reactions critical to life or conversely acting as agents of disease. Mechanical proteins may be grouped into two classes: proteins that act as molecular motors and actively generate force, as well as proteins that are subject to force within their environment.17 Proteins that act as molecular motors include forms such as kinesin, myosin and dynein and are responsible for a host of biological activities like muscle contraction and flagella movement. Proteins subject to force in their environment include elastiomeric proteins. These proteins are specifically engineered to be placed under mechanical tension, serving as molecular springs and adding mechanical strength and elasticity to tissues.18 Such proteins often exhibit a modular structure made of repeated tandem domains. Titin, a key component within striated muscle whose extensibility is critical for balancing forces acting between two sarcomeres during muscle extension and contraction, is a prime example of this type of protein. Upon exposure to force, a modular design means that the applied force may cause a few domains to unfold. This ‘sacrificial’ unfolding maintains the overall integrity of the molecule, protecting it from the destructive effects of force. Unfolding under force also appears to be a design feature of the protein elastin,19, 20 whose lack of structure imparts a high degree of entropic elasticity, allowing connective tissues that have been distended by mechanical force to regain their initial conformation. Elasticity also stems from the reversible nature of this unfolding once the denaturing force is removed. The elastic properties of these proteins, exhibited in their ability to refold accurately after the application of denaturing force, serves a viable biological function. This is in direct contrast with non-mechanical proteins, whose physiological  6  function is not linked intrinsically with force and whose structures do not contain the same force elements that are seemingly intentionally engineered within mechanical proteins. Comparatively studying the folding and unfolding behaviors of these classes of proteins can help elucidate the specific molecular mechanisms that form the basis of mechanical functionality. To date, investigating the mechanical stability of proteins has focused on those mechanical in nature, such as the tenascins, spectrin, and titin. Exploration of nonmechanical proteins has thus far been more limited and resulted in a range of nanomechanical behavior. The small globular RNase protein barnase was one of the first non-mechanical proteins to be examined in terms of its mechanical behavior using single molecule atomic force microscopy (AFM).21 The structure of this enzyme contains none of the structural adaptations for force exhibited by mechanical proteins, and was found to unfold at low applied force. However, studies of the protein T4 lysozyme revealed that this nonmechanical protein shows regular unfolding behavior,22 with unfolding peaks occurring at forces comparable to that of spectrin.23, 24 Being able to pinpoint the molecular determinates of mechanical stability could present the advancement towards the engineering of novel protein based materials, as well as general knowledge of how basic elements within the primary structure of a protein are critical to achieving a high stability fold.  7  Figure 1.2 Mechanical Adaptations of the Muscle Protein Titin A  B Titin Immunoglobulin-like Module Repeat 27  Figure 1.2 The protein titin: a protein demonstrating intrinsic mechanical functionality. The muscle protein titin is found within all striated muscle and is responsible for the elastic response of muscle sarcomeres after muscle extension (Figure A). Titin is a huge modular protein composed primarily of ∼300 tandem repeated immunoglobulin-like domains (Ig domains), fibronectin type III domains (FnIII domains), and random coil PEVK domains. When muscle fibres are stretched, elasticity of the titin protein is partially mediated by the unfolding of Ig-like modules within its structure (Figure 1.2 B). Titin Ig domains exhibit a highly conserved β-sandwich structure. It is thought that Ig domains act to resist the over-stretching of muscle by opposing high mechanical stress and reversibly unfolding under high mechanical loads. Figure 1.2 A used with permission (American Heart Association, Granzier H, Labeit S,Circ Res 2004; 94: 284–95).  1.2 Understanding Protein Folding 1.2.1 Methods of Elucidating Protein Folding Elucidating exactly how and why proteins attain their specific fold has been an exceedingly difficult biological question to answer. A number of different approaches have arisen from the desire to solve details behind folding dynamics, each of which is able to distinctly view but not entirely solve the protein folding dilemma. This speaks to the 8  complex nature of the problem, that protein folding is a dynamic process influenced by a large number of factors and that a simple, black and white solution is not likely. 1.2.1.1 Methods of Determining Folded Structure Knowing the structure of a protein of interest prior to examination of folding details is useful as structural details provides a molecular basis for the folding/unfolding behavior observed. When coupled with simulation techniques such as molecular dynamic simulations (MDS), a clear mechanistic picture of folding or unfolding can be created; obviously, this is made possible only by knowing the folded protein shape. The final structure attained by the protein may be determined utilizing NMR or X-Ray crystallography techniques. As of present, the majority of protein structures contained within the Protein Data bank, a comprehensive compilation of solved protein structures, have been determined by X-Ray Crystallography. This technique involves the crystallization of a purified protein within an appropriate solvent and analysis of the diffraction pattern observed when this protein crystal is subjected to X-Ray radiation. This radiation is diffracted by the 3-D distribution of electrons throughout the protein crystal. High resolution determination of electron densities can be used to determine how atoms are distributed through the uniform crystal. Constructing a basic atomic map for the crystal allows for the overlay of the basic protein shape using the theoretical shapes exhibited by each of the 20 amino acids, giving a three dimensional portrait of how primary structure is arranged into higher order structure. This technique can also determine the structure of proteins bound to ligands, and can thus help elucidate function at a molecular level. Nuclear Magnetic Resonance (NMR) may also be used to determine structural details by aligning nuclei to an external field and observing their subsequent relaxation, which is directly related to the chemical environment in which each NMR active nuclei resides. Because of the sheer size of many proteins, multidimentional NMR experiments, along with techniques such as correlation spectroscopy, Nuclear Overhauser effect spectroscopy and labeling the protein with carbon-13 and nitrogen-15 are used to provide information as to the final structure of the protein.  9  1.2.1.2 Ensemble Methods of Elucidating Protein Folding and Unfolding Studies seeking to elucidate protein folding mechanisms are multiple and all approach the same problem from different angles. Historically, studies have tracked the folding and unfolding behavior of a large ensemble of individual protein molecules. These studies follow observable thermodynamic changes as an ensemble of proteins proceeds from either a folded or unfolded state through the ‘rough energy landscape’ to its reciprocal one. Denaturation, by use of either chemicals or by heat, is the usual means of investigating the unfolding and folding kinetics of an ensemble of proteins in solution. The transition of the protein from folded to unfolded as it is exposed to a greater amount of denaturant is monitored using spectroscopic techniques such as absorbance, fluorescence, circular dichroism, calorimetry or monitoring the loss of activity as an enzyme unfolds. Typically, monitoring changes in absorbance or fluorescence of the protein in solution is used. In the case of chemical denaturation, a solution of protein is exposed to increasing concentrations of a harsh chemical denaturant, such as guanidine hydrochloride or urea. Unfolding or refolding is initiated upon mixing with varying solutions of denaturant in a manner that allows for rapid and complete mixing of the two solutions. This is done using equipment such as a stop-flow apparatus. Monitoring the unfolding and folding kinetics at varying concentrations of denaturant and compiling this data into a chevron plot gives values for both the unfolding and refolding rate constants. Stopped flow denaturation may also be used to determine Cm, the equilibrium concentration of denaturant at which 50% of proteins in solution are unfolded. Thermal denaturation occurs along a similar premise, where an ensemble of soluble proteins is exposed to increasing temperatures until the melting temperature of the protein is exceeded and unfolding occurs. Thermodynamic stability of the protein under study is found using the Tm, or temperature at which 50% of the proteins in solution are unfolded. These methods examine the average behavior of an ensemble of proteins in solution. At equilibrium, soluble proteins will exhibit a probabilistic range of conformations occurring at the bottom of the protein’s energy well, with the majority of proteins conforming to that responsible for the landscape’s absolute minimum. Ensemble techniques provide necessary information as to the average behavior of this spectrum of conformations, the disadvantage to 10  this technique being that subtle information concerning differing behaviors of individual conformers is lost within the averaged ensemble measurement. The resolution of differing folding pathways and the attainment of folding intermediates encountered prior to attaining a final folded state,25 as well as the observation of transient or rare phenomena, is lost by the inherent averaging present in ensemble measurements. 1.2.1.3 Single Molecule Methods of Studying Protein Folding and Unfolding: FRET Several techniques have emerged to investigate protein folding at a single molecule level involving the use of intrinsic fluorescence or force. The most commonly encountered single molecule fluorescence technique is fluorescence resonance energy transfer (FRET). FRET involves the covalent attachment of a donor and acceptor fluorophore molecule to two specific sites on a biomolecule such as protein. A dipole-dipole interaction is induced within proximal donor and acceptor dye molecules resulting in energy transfer, where fluorescence emission is decreased in the donor dye molecule and increased in the acceptor. This efficiency in energy transfer depends on the distance between the donor and acceptor dye molecules R according to 1/(1+(R/Ro)6). An appropriately labeled protein is exposed to denaturing conditions, and conformational changes due to unfolding are monitored by observing resultant changes in fluorescence as the dye molecules change their relative position. FRET, an exceedingly sensitive technique, has the advantage that it may be used either for an ensemble or for a single labeled protein, and may be used either on single molecules free in solution or those affixed to a solid substrate. Fluorescence self quenching is a second, lesser-used fluorescent technique that works along a similar premise. A biomolecule is labeled with two identical fluorescence molecules who exhibit emission self quenching due to intermolecular interactions when the two dyes are within close proximity to one another. As in FRET, this technique may also be used on a single molecule or on the ensemble scale and either on free or affixed molecules. In either technique, unfolding or folding of a protein is usually initiated by either the presence or absence of a chemical denaturant. The free energy pathway followed by the folding or unfolding protein is highly anisotropic in nature; thus, it is likely that a protein denatured chemically does so in a different manner than one denatured by heat or mechanical force.8  11  1.2.2 Single Molecule Force Spectroscopy Force spectroscopy mediates the transition between folded and unfolded states via the application of mechanical force. Mechanical force plays a fundamental role in many biological processes, from cellular motion such as cellular mobility and molecular motors such as kinesin, to the segregation and replication of DNA. The presence of specific mechanical forces is also critical in the processes of cellular adhesion,26 ligand receptor interactions,27 and muscle contraction.28 As previously discussed, the molecular function of mechanical proteins is tailored to either create or respond to mechanical force. As force is encountered within many physiological environments, adaptations that make use of and resist the denaturing effects of such force are commonly encountered within a range of protein folds. 1.2.2.1 Force as a Physiologically Relevant Means of Unfolding Force spectroscopy techniques exploit the biological importance of force and the fact that a large proportion of proteins are not only subject to regular mechanical force but are engineered to do so. Folding and refolding reactions thus occur along a pathway analogous to that occurring within a biological context, whereas the pathway exhibited by a protein exposed to a chemical or thermal denaturing source explores differing regions of the free energy landscape. In a sense, exploring protein dynamics using force spectroscopy could be considered a more ‘true to nature’ technique for some proteins. Experimental comparison between the unfolding pathways exhibited by a protein unfolded either by force or by chemical denaturants supports the conclusion that differing denaturing methods forces the protein down different free energy pathways.8, 21, 29 This was partially established using Φ value analysis to compare the unfolding of I27 caused either chemically or mechanically, where it was found that interactions formed in the A strand of the chemically denatured transitional state did not occur during mechanical unfolding.30 This analysis also showed that the pathway initiated upon mechanical unfolding was not populated during chemical unfolding, and further study of a mutant form of the I27 A strand that destabilizes the native state in chemical unfolding demonstrated no effect on the manner in which the mutant protein mechanically unfolds.8 Using force to denature proteins also has implications when 12  disconcerting the kinetics of an unfolding/folding reaction. The unfolding rate constant may be extrapolated to zero force in principal, as in chemical denaturation. Studying chemical denaturation at a variety of denaturant concentrations assumes that the mechanism that occurs during unfolding is the reciprocal of that which occurs during folding. This assumption is well affirmed for in the case of chemical denaturation, where the presence or absence of denaturant alters the relative stabilities of unfolded and folded species along the energy landscape.31 In contrast to this, refolding that occurs under an absence of force may not be the mechanistic reverse of forcible unfolding, allowing for the investigation of areas of the energy landscape inaccessible by means such as chemical and thermal denaturation. 1.2.2.2 Force Spectroscopy Techniques Several force spectroscopy techniques exist, where the most widely used are optical tweezers, magnetic tweezers and atomic force microscopy. All work by the basic premise where the protein of interest is attached between a fixed surface and a moveable probe in such a way that its biological and mechanical properties are not changed. Force is applied to the system through the moveable probe, which tracks the extension of the molecule of interest through its relative position to the fixed surface. 1.2.2.2.1 Optical Tweezers Optical tweezers utilize an optical trap in order to manipulate a protein-functionalized bead that acts as the system’s probe. Protein studies involve attaching one end of a protein of interest to a microbead made out of a dielectric material such as polystyrene or silica. This microbead acts as the systems probe, where the other end of the molecule is attached to a fixed surface such as a micropipette. The optical trap is formed by focusing a laser onto a diffraction limited spot with a high numerical aperture microscope objective.32 Incident photons colliding with a dielectric material, such as a microbead, near the focal point experience a transfer of momentum from the photon to the bead, which then experiences a restoring three dimensional force. A force felt in the direction of the spatial light gradient is due to the dielectric nature of the microsphere; as it becomes polarized by the optical field, this dipole interacts with the steep light gradient formed near the focal point of the laser,  13  resulting in a force directed along the light gradient of the laser. A scattering force directed along the line of propagation also forms and causes the equilibrium trapping position to lie slightly past the focus. Successful trapping depends on the ability to form a very steep gradient in light, causing the axial component of light to exceed the scattering force pushing the microsphere away from the light source. So long as displacements of the object ‘trapped’ from its equilibrium position are small (~150 nm), force is linearly proportional to displacement; thus, the system acts as a Hookian spring whose spring constant depends on the steepness of the optical gradient. The motion of the bead in solution is monitored by means such as a CCD camera or photodiode to ensure that the bead is trapped in the equilibrium position of the optical trap as well as to track subsequent motion of the bead during application of force to the system. Stretching of the protein molecule is accomplished by movement of the microscope stage, where the three dimensional displacement of the bead from equilibrium within the trap is used to determine force.33 Optical tweezers may be quite dynamic in terms of its experimental setup. This including schemes used to study molecular motor motions of a kinesin molecule moving along substrate-fixed microtubules,34 as well as multiple trap scenarios able to measure the individual base pair steps of RNA polymerase as it translocates down a molecule of DNA.35, 36 Optical tweezers are a powerful single molecule technique, able to measure forces up to 100pN with sub-nanometer spatial resolution and sub-millisecond time resolution.37 These properties have been well exploited in the exploration of molecular motors such as kinesin and polymerases. This does not mean that this technique is without its limitations. This includes concerns such as heating at the optical focal point, optical interference and perturbations affecting the stiffness of the optical gradient, as well as trapping of contaminant particles within impure samples. The largest hindrance towards application of this technique to protein unfolding involves its relatively small range of applied force available (0.1100pN). This makes optical tweezers too ‘weak’ to be applied to many mechanically stable proteins whose unfolding force exceeds the capability of the technique.  14  1.2.2.2.2 Magnetic Tweezers Magnetic tweezers operate by a similar concept to that of optical tweezers. In the case of magnetic tweezers, the microbead composed of a dielectric material is replaced with a super-paramagnetic bead and the optical trap replaced with a trapping chamber formed by the magnetic field gradient induced by permanent magnets. As in optical tweezers, 3-D manipulation of the bead is possible by correlating motion of the external magnets, or current if electromagnets are used, with that of the magnetic bead. Magnetic tweezers allow for the manipulation of single molecules with forces of 0.01-100pN. However, magnetic tweezers lack the versatility, temporal and spatial resolution of both optical tweezers and AFM. Magnetic tweezers has proven a useful technique in the study of nucleic acid enzymes38 and the rotary motor of F1 ATPase39 due to the ability to impose rotational motion onto a system. 1.2.2.3 The Atomic Force Microscope 1.2.2.3.1 Origins and Technological Advancements of the Atomic Force Microscope The atomic force microscope (AFM) was originally developed in 1986 as a means of imaging nonconductive surface with extremely high resolution. The developmental roots of the AFM lies in scanning tunneling microscopy (STM), an imaging technique developed in 1981 that uses the electron tunneling current formed between a sharp tip and a conductive surface in order to create an image of the surface scanned by the probe. The STM technique allowed for imaging at an unprecedented scale, showing atomic detail on a surface while bypassing the diffraction limit in a manner that has higher sensitivity and is less energy intensive than electron microscopy. The discovery of the STM was a major advance in imaging techniques, earning Gerd Binnig and Heinrich Rohrer the 1986 Nobel Prize in physics for their invention of the technique. The desire to extend the high resolution of the STM technique to non-conductive materials resulted in the invention of the AFM technique in 1986 by Gerd Binning, Calvin Quate, and Christopher Gerber. This preliminary AFM had some of the features of a modern AFM, with the sample connected to a piezoelectric stage responsible for modulating the interaction between the sample surface and the AFM tip, where, in ‘contact mode’, surface features are revealed upon detection of cantilever  15  deflection. The detection of cantilever deflection was accomplished using a STM tunneling tip in contact with the conductive surface of the AFM tip, where deflection due to surface features detected upon changes in tunneling current between the STM tip and AFM cantilever.40 The newly minted AFM system, which at first was really a technical improvement on existing STM technology, had the atomic imaging resolution of the STM, with Angstrom vertical resolution,41 without the need for a semiconductor substrate. The AFM has undergone a number of technical and procedural iterations that has allowed it to become a powerful instrument useful in a variety of applications above that of simply producing a high resolution image. Technical advancements include the invention of micro fabricated cantilevers and the use of an optical detection system42 have delineated AFM from STM as well as offering increased functional utility. Conformation of the fundamental accuracy of data obtained using the AFM was also an early worry, with the onus being on whether the AFM was actually making measurements from a single molecule. Substantiation of the validity of information obtained by AFM was mostly agreed upon within the scientific community by the middle of the 90’s.43 The AFM has evolved into a technique capable of high spatial resolution, dependent on the sharpness of the probe tip used, as well as high force resolution due to the small spring constant of the cantilever utilized. The AFM as a Widely Utilized Imaging Technique The applicability of the AFM in imaging biological samples was quickly realized and represented a chance to investigate samples that were previously irresolvable by existing methods. The first image of a biological sample produced using AFM was done on bulk crystals of amino acids.44 Later, in-liquid AFM methods were introduced to image solution state samples,45, 46 a critical development allowing for such advances as the first topography of membrane proteins with molecular resolution, showing the dissection of the top layer of the double-layered gap assembly.47 Further advancements in the technological evolution of the AFM included the introduction of non-destructive vibrational48 and tapping modes.49 The development of low or non-contact procedures was a critical improvement upon existing techniques due to the softness of most biological samples, where application of undue force 16  resulted in sample destruction. The utility of the AFM as an imaging technique has been well demonstrated, from the resolution of individual beta-turns on the surface of the membrane protein porin OmpF50, to the development of in vivo cellular imaging methods used for fungal,51 bacterial,52 and mammalian cells.53 Development of the AFM as a Force Spectroscopy System The development of the AFM beyond that of a high resolution imaging technique occurred relatively early in the timeline of its development, a logical turn that realizes the applicability of AFM to a number of different environments and states as well as the utility of using mechanical force to probe biological systems. In the first case, AFM may be used on a number of sample conditions, from aqueous to vacuum and at a large range of temperatures and pH. The AFM was demonstrated to work within aqueous systems in 1987,46 a critical step towards the emergence of the AFM as a force spectroscopy rather than imaging system.27 The high sensitivity at which the AFM applies force to a system of interest makes it applicable to the direct measurement of force at the microscale. Thus, the AFM tip transformed from a passive tip scanning a surface to be imaged to an active participant within the system being probed. This tip acts as the probe within the system, able to detect extremely localized force from few picoNewtons (pN) up to a few hundred picoNewtons. Such force is well beyond the reach of other force spectroscopy techniques. The applicability of the AFM force spectroscopy approach extends to a huge number of biological systems. This includes the study of bimolecular interactions,27, 54, 55 probing the viscoelastic properties of whole cells,56-58 the study of biomembrane proteins59-61 and surface receptors.59, 62, 63 Single Protein Molecule Iterations of the Atomic Force Microscope The first study to use the AFM to study the unfolding and refolding dynamics of a single molecule of protein, the predecessor of the research presented here, occurred in 1997. This study used AFM to induce unfolding of multiple domains of titin, the giant sarcomeric protein present within striated muscle fibres.59 This was the first investigation to directly study the mechanical properties of a single molecule of protein whose functionality was known to be highly mechanical. It was also the first introduction of an experimental setup  17  mirroring the one used subsequently within research presented here, where a single molecule of protein is attached to the cantilever of an AFM, whose deflection under forced unfolding monitors the conformational changes in the protein as it unfolds. These force extension cycles yielded a spectrum mapping the unfolding of the protein in a ‘sawtooth pattern’ formation in which the nanomechanical properties of the domains subject to unfolding force are encoded. This allowed for the elucidation of the mechanical stability of titin, as well as the observation of folding characteristics such as the protein’s intrinsic elasticity and ability to refold after unfolding. The capacity of AFM to provide detailed insight into the folding mechanism of proteins, as well as the advantage of using force over chemical or thermal denaturants, fostered the growth and development of AFM into the valuable single molecule force spectroscopy technique that it is today. Since 1997, the AFM has also been utilized as a method of detailing unique DNA64-67 and polysaccaride8, 59, 68 mechanisms, though it has arguably enjoyed the most success in its application towards defining protein dynamics. To this end, the AFM has been used in combination with techniques such as molecular dynamics simulation and molecular biology to help elucidate information about the contribution of specific structural motifs to stability as well as the elucidation of entire folding energy landscapes. Thus, the modern AFM has become a powerful tool in investigating the mechanical properties of proteins at the single molecule level.  18  1.3. The Atomic Force Microscope 1.3.1 Overview of the Atomic Force Microscope Figure 1.3 Schematic of the AFM Instrument  A  Laser  B  Photodiode Detector Cantilever  F  Silicon nitride tip  Glass Coverslip  Piezoelectric Positioner  Figure 1.3 Overview of the AFM instrument. The protein under investigation is adsorbed onto a clean glass coverslip which is subsequently mounted onto a piezoelectric positioner. A single molecule of protein is attached between this substrate and a silicon nitride tip via upward movement of the piezoelectric positioner, which presses the tip into the adsorbed protein layer. Force is applied to the attached protein by downward movement of the piezoelectric positioner, causing an opposing entropic restoring force originating from the protein which forces the downwards deflection of the cantilever. Cantilever motion is observed by the reflection of a laser focused tightly onto the cantilever end; as the cantilever is deflected downwards, the laser spot focused onto a laterally divided photodiode also deflects, creating a discernable output signal relative to the force extension of the protein.  The atomic force microscope setup utilized within all subsequent experiments can be loosely divided into three components: a piezoelectric positioner fitted with a glass coverslip onto which the protein in solution is adsorbed, the actual AFM head containing a silicon nitride cantilever and laser detection system, and the control system monitoring the position of the piezoelectric head and the subsequent deflection of the cantilever. The particular setup used is based upon those described previously.8, 17, 59 These systems are intrinsically linked; 19  the protein of interest is adsorbed onto a glass coverslip, which is placed on the piezoelectric positioner manipulated by the feedback system. The AFM head contains the cantilever and optical detection system; it fits over the coverslip with the thin layer of adsorbed protein, and force is applied to a single molecule of protein attached between the cantilever and glass substrate via movement of the piezoelectric positioner. Force induced conformational changes of the protein causes the cantilever to deflect, the magnitude of which is detected by monitoring the motion of the laser spot reflected off the cantilever tip. The control system includes the data acquisition system, the piezoelectric positioner control, an optional oscilloscope and computer software responsible for setting the movement of the piezoelectric positioner and integrating this motion with the deflection of the cantilever. 1.3.2  The Piezoelectric Positioner and Glass Coverslip The source of applied force within the experimental setup utilized is the piezoelectric  positioner. This positioner is capable of ultra precise movement, with spatial resolution along the z-axis approaching the truly ‘atomic’ scale. Two types of positioner were utilized, one capable of movement in the z-plane and one capable of movement in all three directions. Force is applied to the single molecule of protein between the piezoelectric positioner and cantilever via movement of the positioner in the z-direction. The precision of the piezoelectric positioner is important within single molecule studies when the scale of the experiment is considered. Elongation of single molecules occurs at the nanometer scale and relevant conformational changes occur within the picoNewton range, making it necessary that instrumentation operates precisely within this exceedingly small, narrow range. This makes the piezoelectric positioner uniquely adapted towards single molecule studies, with its nanometer scale spatial precision and ability to move in a z-plane with accuracy and precision. Mounted on the moveable piezoelectric positioner stage is a solid substrate onto which the protein under study is attached. In terms of the research presented here, this solid substrate was a simple glass coverslip. Cleanliness as well as surface hydrophilicity of this surface was assured by boiling and subsequent storage of the coverslip within a sulfochromic solution. Prior to experimental use, coverslips were rinsed thoroughly using deionized water 20  within ~12 hours of use; coverslips older than this were noted to lose their hydrophilicity and were not optimal for the adsorption of protein onto their surface. The protein of interest was introduced by pipetting a known concentration of the soluble protein in phosphate buffered saline (PBS) solution and allowing it to adsorb to the glass surface for at least 15 minutes prior to further experimentation. For the purposes of the experiments within this document, protein adsorption occurs non-specifically via physisorption to form a layer 20-50 nm thick.69 Functionalization of the surface was deemed unnecessary for a number of reasons. Firstly, the concentration of the protein within solution was optimized such that concentrations were not so low as to make it impossible to attach a molecule of protein between the surface and cantilever, but not so high as to make single molecule experiments impossible. Optimal protein surface concentrations were within the milliMolar range. Successful single molecule experiments depend on cleanliness, where reliable data rests on the ability to confidently determine whether the experiment is indeed preformed on a single molecule of protein. To this end, non-specific attachments between the substrate and cantilever were greatly decreased if excess buffer containing non-bound protein was pipetted from the glass surface after allowing a sufficient adsorption time to pass and introducing fresh buffer in which to perform pulling experiments. Lastly, two cysteine residues were engineered on the Cterminus of the protein in question, knowing that this addition allows for the covalent attachment of proteins onto a gold surface59 and seems to increase the rate of adsorption onto hydrophilic surfaces. Hydrophilic glass coverslips provide a cheap method of supporting the thin layer of adsorbed protein molecules with which to study; however, multiple other schemes aim to increase the successful adsorption of protein as well as to increase the specificity with which proteins of interest adsorb. Evaporated gold surfaces increase adsorption through covalent bonding between the gold surface and sulfur groups contained within cysteine residues engineered into the protein of interest. Surfaces such as cleaved mica ensure a molecularly smooth substrate onto which proteins adsorb, reducing possible effects from large surface aberrations found in surfaces such as glass. Protein adsorption is a nonspecific process in that all molecules with an affinity for the surface will also adsorb, affecting the precision of the subsequent experiment. Within an experimental setup utilizing an unfunctionalized  21  surface such as glass, this specificity is guaranteed by ensuring the purity of the protein solution introduced to the slide. However, aberrations caused by surface contamination can make data analysis difficult, and functionalization of the surface such that only the protein of interest will attach help to decrease the likelihood of contamination. Functionalization of the substrate surface also allows for a large degree of control over factors such as surface concentration, and gives the ability to pattern the surface. One common surface functionalization scheme involves the adsorption of streptadivin onto a surface such as glass or cleaved mica. Streptadivin binds the small peptide biotin with high affinity, and it is this affinity that is used advantageously with the tagging of biotin to the protein of interest. Molecules tagged with biotin will be bound with overwhelming preference over untagged contaminants, greatly increasing the specificity of subsequent experiments. All of these methods may be advantageous in offering a higher degree of specificity or protein adsorption, but are more costly and require a great deal more preparation than simply using glass substrates. Careful sample preparation, as well as proactive data analysis, meant cleaned glass coverslips were more than adequate for evaluating the proteins presented within this work. 1.3.3  The AFM Head The AFM head is the heart of the AFM system. It consists of a removable clear  quartz stage in which the silicon nitride cantilever and syringe containing buffer are loaded as well as the laser based optical detection system. The AFM cantilever is responsible for the initial detection of force induced conformation changes of the protein molecule as it is unfolded or allowed to refold. The properties of the cantilever, specifically its stiffness, determines the force range in which the AFM may operate; the relative stiffness of the cantilevers in AFM use compared to optical or magnetic tweezers impart the high force range critical when unfolding proteins. The AFM cantilever is composed of two parts; the actual cantilever whose stiffness determines the spring constant of the system as well as a tip or probe with nanoscale sharpness that attaches to the molecule under study. AFM cantilevers are quantified according to their width, length and thickness, properties that affects the resultant spring constant as well as the resonance 22  frequency important to other types of AFM use. AFM cantilevers are manufactured out of silicon or silicon nitride and can be coated with metals such as aluminum or gold in order to impart improved conductivity on the tip or to improve its reflectivity. In the case of this study, the cantilever used was made of silicon nitride with a reflective gold coating, with a tip made only of silicon nitride. The basic AFM setup is incredibly versatile, and may be employed in anything from imaging to nanolithography to force spectroscopy; this diversity is reflected in the range of AFM tips available, each with different specific functionality. Tips are micro fabricated in a variety of shapes and sizes, from relatively large pyramidal shapes down to carbon nanotubes responsible for extremely high resolution imaging. In order to be useful within single molecule force experiments, tips do not need conductive coatings or the sharpness that conductive or imaging techniques require. A typical radius of curvature is ~50 nm, much larger than the size of a typical protein (~5 nm), its use over that of a blunter tip ensures the attachment of a single molecule of protein. In a setup involving a nonfunctionalized tip, the protein of interest attaches to the tip via non-specific interactions, which poses the same problems for specificity as adsorption of the protein to the substrate. Tips functionalized in a manner similar to that of the substrate have arisen to combat this. Functionalization pertinent to single molecule studies usually occurs through attachment of an amine to a silane or ester functionalized tip. A long polymer such as polyethylene glycol (PEG) is then attached to the amine, and finally a biological molecule made to attach to the protein of interest is attached to the long PEG spacer. These biological molecules could be streptadivin-biotin pair,70, 71 or they could be an antibody pair.62, 72, 73 Tip functionalization does prevent the nonspecific interactions that occur between the cantilever tip and any contaminants that may lie on the surface of the substrate. Like substrate functionalization, these methods involve a number of air-reactive steps that greatly increase the complexity of the experimental setup, and are not necessary so long as solution purity is ensured and data is carefully analyzed. Within this context, unfunctionalized tips made of silicon nitride were more than adequate to supply meaningful results.  23  1.3.4  Cantilever Calibration and the Equipartition Theorem In order to determine the relationship between the deflection of the cantilever and the  actual amplitude of the restoring force causing it, the spring constant and the ratio of the photodiode output voltage to cantilever deflection must be elucidated. The spring constant is critical to ascribing force to the motions of the cantilever, which behaves as a Hookian spring. Based on this relationship, the restoring force placed on the cantilever by the protein depends on both its spring constant k and displacement of the cantilever along the z axis ∆zc: F=-k zc  (1.1)  Determination of the spring constant k is most commonly done using the thermal method. In this, the cantilever is modeled as a simple harmonic oscillator being oscillated by environmental thermal noise. The magnitude of the oscillation frequency and amplitude depends both on temperature as well as the same mechanical properties of the cantilever that determine the spring constant. This relationship is known from the equipartition theorem:74  1 1 ks ∆x 2 = k Bt 2 2  (1.2)  In this, the mean-square displacement noise ∆x is determined from the free oscillation of the cantilever suspended in solution. Cantilever fluctuation data is transformed using a fast Fourier transform into its corresponding power spectrum. The thermal contribution to ∆x is determined by integrating over this power density spectrum as only thermal motions are within the resonance range of the cantilever.74 Calculation of the spring constant using this method results in a relatively large error (~20%),74 but has the advantage that it requires no specialized equipment and is accomplished easily prior to the beginning of an experiment. The experimental spring constant was relatively high and varied from 50-80 pN/nm, ensuring that the dominant elastic element within the system is the protein under consideration.  24  1.3.5  The Optical Detection System Detection of cantilever deflection as it experiences the restoring force as a protein is  stretched is done using an optical laser system. In this, laser light is focused on the tip of the cantilever. This light is reflected back upon a photodiode detector split into an upper and lower segment. The split photodiode converts incident light into voltage and reports the voltage difference between the two sections. At equilibrium, when the cantilever is unbent, the reflected laser spot is calibrated such that the output of both photodiodes is the same and the spot is balanced evenly between each. When the cantilever begins to bend, the deflected laser spot drops, and the voltage is incurred predominately from the lower photodiode. The change in voltage difference ∆V observed as the spot position changes may be related to the cantilever displacement ∆zc by the ratio ∆V/∆zc. This ratio is determined by bringing the cantilever tip into contact with the substrate, causing the cantilever to deflect, and subsequently retracting it. The deflection and subsequent retraction of the tip from the surface produces a linear voltage vs. deflection response. The amount that the cantilever has deflected is equal to the displacement of the piezoelectric, which can be accurately determined from the strain gauge position sensor within the previously calibrated positioner. The slope of this displacement gives ∆V/∆zc, which is used to convert the voltage output of the photodiode V into cantilever deflection (in nm), allowing force to be calculated from photodiode output according to Equation 1.1. The extension of the molecule Ex may be calculated from the movement of the piezoelectric positioner ∆zp and the subsequent deflection of the cantilever ∆zc according to: Ex= ∆zp- ∆zc 1.3.6  (1.3)  Piezoelectric Positioner Driver and Output Software The final pieces of the AFM setup involve the driver responsible for accurate motion  of the positioner as well as computer software integrated with this driver. This integrated computer-driver system allows for both the manual as well as automated control of the piezoelectric positioner. The IGOR computer software also acquires force data from the piezoelectric positioner as well as voltage output from the photodiode to form the force (pN) 25  versus extension (nm) curve that is the experimental end point in extending a single molecule of protein. An oscilloscope can also be used to monitor the real-time position of the cantilever relative to the sample surface. 1.3.7  Experimentally Pulling a Single Protein Molecule Prior to beginning a single molecule experiment using the AFM, a single molecule of  protein must be attached between the thin layer adsorbed onto the glass and the cantilever tip. This is done by pressing the tip into the protein layer and retracting it, picking up a single molecule of protein at random along its length. Subsequent downward motion of the piezoelectric positioner places a mechanical force on the protein constrained between the tip and the substrate. This results in the unfolding of the protein if its mechanical stability is exceeded by the force applied. The motion of the piezoelectric positioner, and the subsequent application of force, may occur in either a constant speed (speed clamp) or constant force (force clamp) manner. 1.3.7.1 Force Clamp Experiments In constant force experiments, the mechanical force applied by the piezoelectric positioner is kept constant through the experiment by varying the rate at which the positioner moves. This is done using a feedback loop between the photodiode and piezoelectric positioner where the length of the molecule is adjusted to maintain a constant cantilever deflection. This technique monitors the length changes in the molecule with respect to time to determine the unfolding pathway taken as the molecule is subject to a constant mechanical unfolding force. This method allows for the clear determination of kinetic ‘steps’ taken by the protein as it unfolds under force. However, it is also prone to error from cantilever drift. Implementing a ‘force ramp’ type of ‘constant force’ experiment has arisen to deal with cantilever drift intrinsic to all single molecule AFM experiments. 1.3.7.1 Constant Speed Experiments The more commonly used scenario, and the setup utilized here, is of the constant speed or ‘speed clamp’ type. In this setting, a single molecule of protein is picked up along 26  its contour and forms the elastic element in the mechanical circuit between tip and surface. The piezoelectric positioner retracts at a constant rate. As this rate increases the extension of the molecule, the force applied to the molecule also increases until the mechanical stability of the protein is exceeded and it unfolds. Unfolding of the protein results in a sudden increase in the length of the protein, which allows the cantilever to return close to its undeflected position. Unfolding of multiple domains occurs as the piezoelectric positioner is continually retracted until the protein is entirely unfolded and detaches from either the surface or tip. To investigate the refolding of a molecule, the piezoelectric positioner is moved upwards back towards the original, unstretched position after the unfolding of domains within the polyprotein attached between tip and surface. By removing force from the system in this manner, elastic proteins may refold into their original conformation and be re-unfolded in the manner described. Force changes observed from cantilever deflection compared to the extension of the piezoelectric positioner results in a force extension curve that provides a wealth of information concerning the nanomechanical properties of the protein under investigation. The constant speed method may also be varied with refolding or speed dependent experiments in order to elucidate refolding and unfolding kinetic parameters. 1.4 Investigating the anomechanical Fingerprint of Mechanical Unfolding 1.4.1 General Features of the Force Extension Curve The general features of the force extension curve arise from the behavior of the protein as it is subject to increasing mechanical forces and may be separated into two portions. The extension phase of the force spectrum is due to the increasing force applied to the protein as it is continually stretched by the motion of the piezoelectric positioner. Once the mechanical stability of the protein is exceeded, it unfolds typically in an all or none fashion. Unfolding results in a sudden increase in the contour length of the protein, which causes the cantilever to return to a much less deflected position. Proteins most appropriate for study using single molecule AFM are typically composed of multiple individually folded domains, which means that this process is repeated following the unfolding of one domain. Continual extension so that all domains are eventually unfolded gives peaks which form the ‘sawtooth pattern’ typical to AFM studies, with the last peak corresponding to the 27  detachment of the molecule from either the tip or substrate. Unfolding of domains is typically arranged in a hierarchy, with unfolding events occurring at high extension corresponding to unfolding of domains with the greatest values of mechanical stability.  28  Figure 1.4 Unfolding a Modular Protein Results in a Representative Sawtooth Pattern Piezoelectric Positioner  Cantilever  D  C  B ∆zp  ∆zc  A  F=  k BT  1  x 1 − P  4  LC   ∆Lc  −2  1 x   − + 4 LC     D  A  C  200 pN  B  50 nm Figure 1.4. The sawtooth unfolding pattern of a modular polyprotein unfolded using single molecule AFM in distance ramp or constant speed mode. Proteins adsorbed onto a glass surface are picked up at random along their contour by the cantilever tip. Initially, both the piezoelectric positioner and cantilever tip are within a small distance of one another, and the polyprotein chain linking the two is relaxed (position A). Retraction of the piezoelectric positioner ∆zp applies a force F along the protein chain. This applied force is initially resisted by an entropic restoring force that causes the downward deflection of the cantilever (position B, ∆zc). If the applied force exceeds the mechanical stability of one of the domains within the polyprotein, the domain unfolds resulting in a peak on the sawtoothpatterned force extension curve. The sudden unfolding of a single domain results in a sudden increase in the contour length of the polyprotein, decreasing the deflecting force acting upon the cantilever, allowing it to return close to its initial position (point C). Subsequent retraction of the piezoelectric positioner repeats this process, resulting in a number of unfolding peaks within the sawtooth pattern. As pickup occurs nonspecifically along a polyprotein chain, the number of unfolding events observed within a single force extension curve varies, corresponding to the number of domains contained within the stretched polyprotein. Unfolding events may be fit using a Worm-like Chain model of polymer elasticity (red fits, Equation 1.4), which details the experimental contour length increment (∆Lc) of each domain. The last peak within the force extension curve corresponds to the detachment of the unfolded polyprotein from either the surface or tip.  29  1.4.2 Improving Features of the Force Extension Curve Using Polyprotein Constructs Information presented within the force extension curve regarding protein unfolding is made clearer and exponentially easier to interpret if multiple unfolding events are presented. Proteins such as titin that are composed of multiple individually folded domains are naturally attuned towards single molecule studies. Investigation of a single individually folded domain is improved by engineering this domain into a polyprotein construct composed of either multiple repeats of the protein in question or alongside a protein used as a nanomechanical fingerprint within force extension curves. Multiplicity is advantageous for a number of reasons. The low extension portion of the force extension curve is typically obscured by non-specific interactions that occur between the tip and surface, obscuring any unfolding events that would occur in the region. Unfolding of a polyprotein results in multiple unfolding events, some of which will occur at higher extension than that of a single domain. The sawtooth pattern created by the unfolding of a polyprotein is distinct and easily predicted from a theoretical contour length increment, and because of this single molecule events occurring are easily identifiable. As well, adsorption of a domain to either the tip or to the surface usually results in some degree of unfolding, making it necessary to include ‘sacrificial domains’ along the length of the protein. Some degree of homogeneity is useful in this case, as pickup occurs in a random manner, making it impossible to accurately assign specific domains to their corresponding unfolding events and to predict which domains will act sacrificially. Use of repetitious polyproteins demonstrates the stochastic nature of protein unfolding, in that unfolding behavior of a single protein domain may vary slightly if its unfolding behavior is compared over a large number of extensions. This is a fundamental lesson to be learned in protein unfolding, and one easily illustrated when linking multiple unfolding events into a single extension curve. Most importantly in terms of single molecule experiments is the fact that the use of polyproteins allows for the unambiguous determination of single molecule unfolding events through identification of a well defined, repeating sawtooth pattern. Experiments that do not exhibit reproducibly spaced unfolding events within their force extension curve, similar to that observed in Figure 1.4, are not single molecule unfolding events. If a homogenous polyprotein is utilized, data acquisition is sped considerably as multiple unfolding events may be analyzed by pulling one molecule.  30  Although the use of heterogeneous polyproteins such as titin reduces the specificity of assigning specific unfolding events to their corresponding domains, the use of polyproteins is critical in assuring a well defined single molecule experiment. 1.4.3  Entropic Restoring Forces and the Worm Like Chain Model As a protein molecule is extended, a restoring force is generated to resist the forced  extension, causing proportional deflection of the cantilever. The resistance of the protein to extension is entropic in nature, as a folded protein attempts to maximize its conformational entropy by remaining so. Proteins under force act as entropic springs, where a resisting force opposing extension arises from this attempt to maximize entropy under the drive of thermal fluctuations.75 Entropic elasticity arises as a general property of flexible polymers such as proteins, and can be formally described using the Worm-like Chain (WLC) model of polymer elasticity. Within the WLC model, polymers are envisioned as an entropic rod continuously flexible along its length. This rod is divided into equal flexible segments that align with one another and, at rest, attain a smoothly curving conformation. In terms of single molecule protein studies, the iteration of the WLC model that fits the entropic elasticity demonstrated is not an exact solution. Instead, an interpretation made by Bustamente et. al. (1995) and Marko and Siggia (1994) describing the elasticity of double stranded DNA is extensively used to describe the extension of proteins. The applicability of this approximation to both systems is possible as they are both relatively stiff rod polymers; this form of the WLC poorly describes more flexible polymers. Within this iteration of the WLC, the restoring force F due to an extension x depends only on two changeable parameters; the persistence length (P) and contour length (LC).76, 77  k T 1 x F = B  1 − P  4  LC   −2  1 x   − + 4 LC     (1.4)  1.4.3.1 Persistence Length as a Measure of Polymer Stiffness The persist length P is defined as the shortest length over which a polymer segment remains rigid. The persistence length is a mechanical property describing polymer stiffness, 31  with stiffer polymers demonstrating higher values of persist length than more flexible polymers. Values of persistence length are determined by fitting to experimental data, and are typically on the order of 0.5 nm. 1.4.3.2 Contour Length and the Experimental Contour Length Increment The maximum end-to-end distance of a polymer is defined as its contour length. In the case of single molecule protein experiments, this is the length of the peptide when it is fully unfolded and can be easily calculated by multiplying the number of amino acids within an individually folded domain by the average length of an amino acid (0.36 nm). In the force spectrum of a protein composed of multiple individually folded domains, the contour length increment determines the spacing between Worm-like chain fits of consecutive unfolding events (Figure 1.4, red lines). The contour length increment ∆Lc is detailed by the number of amino acids within the domain architecture that are exposed by unfolding (Figure 1.5). The region that these amino acids detailed has also been termed the ‘force hidden’8 region of the protein, since the strength of the bonding that occurs between these residues ensures that without force these regions of protein remain tightly folded. Unaccounted changes between contour length and ∆Lc may be accounted for in a number of ways. For a multidomain polyprotein such as titin, the difference between the contour length containing all residues annoted within the protein primary structure and the experimental contour length increment could signal the presence of linker sequences that do not participate in folding. The conformation of the folded protein is accounted for within the contour length increment by a decrease in the contour length by the distance with which the N and C terminus is separated and can be estimated so long as a crystal structure of the protein is available (Figure 1.5). Covalent disulfide bonds that occur wherever cysteine residues are within proximity of one another are too strong to be disrupted by the forces generated by the AFM, trapping any residues occurring between the bonding residues. Unexpected changes in contour length increment between sequential peaks in the sawtooth pattern could also show the formation of unexpected secondary structure as the protein unfolds. For example, observed ‘skip’ peaks that occur in a small number of unfolding events exhibited larger than expected ∆Lc within FnIII domains from tenascin are caused by the formation of a ‘superfold’ structure composed of two consecutive domains.8, 75 32  Figure 1.5 Representations of the Contour Length and ∆Lc  F  Lf ∆Lc  F  Lu Lu=(No. of amino acids)(0.36 nm/a.a) ∆Lc=Lu-Lf  Figure 1.5 Representation of the contour length (Lu) and contour length increment (∆Lc). The contour length is the maximum end to end length of an unfolded protein, and is found by multiplying the number of amino acids within the protein sequence by the average length of an amino acid (0.36 nm/a.a) as shown. The contour length increment details the number of amino acids participating in folding and differs from the contour length increment by the length of the folded protein Lf. Values of ∆Lc are experimentally determined by fitting unfolding events using the Worm-like chain model of elasticity (Figure 1.4, red fit lines).  1.4.4  Mechanical Unfolding and Unfolding Kinetics of Proteins The unfolding force of proteins is determined from the local maximum attained by a  single unfolding event within the sawtooth pattern exhibited by an unfolding protein. This value relates to the mechanical strength of the domain being unfolded. Typically, the unfolding force value is reported from an average made over a large number of unfolding events from a number of different single molecules. As unfolding is stochastic in nature, averaging over a large number of events increases the confidence in which an unfolding value can be assigned to a specific protein. This lessens the effects that may occur if a single molecule with anomalous stability is examined, as well as variability caused by experimental drift, without obscuring events such as transitional state formation that are lost within ensemble methods. The mechanical stability of a protein, elucidated by this distribution, is intrinsically linked to the free energy landscape of the transition being probed, giving valuable information about the inherent energetic and kinetics of protein unfolding and folding. 33  1.4.4.1 Exploring the Kinetics of the Folded to Unfolded Transition Under Force Prior to the addition of force, the protein molecule trapped between the substrate and cantilever is in a folded conformation in which its free energy is at a minimum. In terms of the rough energy landscape, the protein lies either at the absolute minima or in a conformation close to this minimum. Unfolding is thermodynamically unfavorable due to the steep gradient of the activation energy lying between folded conformations and higher energy denatured conformations. Unfolding is a probabilistic event, meaning that it is possible that a native protein can convert through this energy barrier to a higher energy denatured conformation; for a thermodynamically stable protein this transition is unlikely and not observable within the AFM timescale. Proteins that unfold spontaneously do so because of environmental thermal energy. Thermal energy continues to contribute to protein unfolding under force, and becomes apparent when a large number of unfolding events are collected in a histogram of unfolding forces. This distribution is sloped, where lower force events occur with a greater probability than those at higher force. Though the energy landscape contains a number of local maxima corresponding to the formation of transitional states lying between the folded and unfolded state, unfolding may be portrayed as a transition from a low energy folded conformation across an energy barrier to a higher energy unfolded conformation without significant population of transitional conformations. Proteins are thus assumed to fold and unfold in a two state process according to conventional state transition theory.78  34  Figure 1.6 The Free Energy Representation of Two State Folding/Unfolding  N  U  T  ∆GT-U  ∆GT-N  Free Energy  ∆GU-N  ∆xu  ∆xf Reaction Coordinate  Figure 1.6 The free energy depiction of two state protein folding/unfolding. The folded protein state N is separated from the unfolded state U by a high free energy transition state T. Unfolding is determined by the free energy barrier ∆GT-F, where ∆xu is the distance from folded to transitional state. Folding is determined by the free energy barrier ∆GT-U, where ∆xf is the distance from the unfolded to transition state along the reaction coordinate. The thermodynamic stability of a protein is determined by the energetic difference between the folded and unfolded states, ∆GU-N.  According to this two state model, the folding rate constant kf for the above conversion may be given as:  kf = κ  k BT − e h  ∆GT − D k BT  (1.5)  The unfolding rate constant ku is given as: kT − ku = κ B e h  ∆GT − k BT  (1.6)  Where κ is the transmission coefficient, kB is the Boltzmann constant, T is temperature, h is the Planck constant, and ∆GT-D and ∆GT- the difference in free energy between the transition to denatured and transition to native state respectively. 35  Proteins made to unfold under force do so because the application of force tilts the energetic positions of the native and unfolded state relative to one another. Given sufficient force, the folded conformation becomes energetically less stable than the denatured conformation. Because the amount to which the native conformation is destabilized by force is relatively greater than that of the denatured conformation, the energy barrier between the unfolded and folded states is increased, and the protein remains unfolded in the presence of continued force. Figure 1.7 Energetic Changes Caused by Application of Force N  T  U  Free Energy Reaction Coordinate Figure 1.7 Energy landscape changes upon application of force. The blue curve describes the folding/unfolding energy landscape of a protein unaffected by mechanical force, where the red curve inscribes the energy landscape of a protein exposed to mechanical force. This mechanical force acts to destabilize the folded N, transition T and unfolded U states by varying amounts. The largest destabilizing effect acts upon the native N state; because of this, the barrier ∆GT-N is decreased and the probability of observing unfolding is increased. Because the unfolded state U is comparatively unaffected by the application of mechanical force, the folding barrier ∆GT-U is increased, and unfolded proteins tend to remain so in the continued presence of mechanical force.  Such a drastic change in the protein landscape of the protein has obvious implications for the unfolding and folding rates of the protein. The manner in which force affects the lifetime of a bond was first elucidated by Bell (1978), and further interpreted by Evans and Richie in 1997. This model demonstrates how the intrinsic rate constants of folding and unfolding are affected by force from basic classical mechanics.79-81 36  kT − k f (F ) = κ B e h k T − ku ( F ) = κ B e h  ∆GT −U + F∆x f k BT  ∆GT − − F∆x u k BT  = β 0e  = α 0e  −  F∆x f k BT  (1.7) F∆x u k BT  (1.6)  Thus, folding and unfolding rates become dependent on the force F applied. Kinetically, unfolding is observed when an unfolding force Fu increases the unfolding rate by decreasing the activation barrier by Fu∆xu while also decreasing the folding rate. Also given in this model is the folding and unfolding rate constant at zero force (βo and αo respectively). It is assumed that both ∆xf and ∆xu are independent of force, an assumption that may not hold if the energy landscape is unduly curved and would necessitate the use of another kinetic model.  37  1.4.4.2 The on-Equilibrium ature of Unfolding Under Force Using force to unfold many polymers, such as the secondary structures of RNA and DNA, results in a transition from a native to denatured state that is easily reversible. This reversibility occurs so long as the rate of unfolding is approximately equal to the rate of folding. Equilibrium refolding of the molecule demonstrates a reciprocal force extension curve to that found during the unfolding of the molecule. Reversibility is a signal that the unfolding/refolding reaction probed is at equilibrium. The secondary interactions that occur within the structure of RNA do so in an additive and independent manner, thus making it possible to reversibly refold after being forcibly unfolded. Certain proteins are also exhibit equilibrium folding and unfolding. These proteins exhibit refolding traces that are the mirror of those found as the protein is unfolded. These elastiomeric proteins, including elastin and domains PEVK and N2B of cardiac titin, act as perfect elastic elements able to store and restore elastic energy with 100% efficiency. This contrasts with most proteins, in which secondary structure formation and stability depends on their larger tertiary context and unfolding tends to happen in an all-or-none fashion. As well, unfolding of the molecule using force spectroscopy typically occurs much faster than the rate of equilibration, and the unfolding transition initiated by mechanical force occurs far from equilibrium. This is apparent in the hysteresis that occurs between the unfolding and refolding force extension traces as well as the typical asymmetry that occurs between values of ∆xu and ∆xf.24 Non-equilibrium unfolding and refolding is thought to dissipate elastic energy into heat as a method of absorbing force in a type of ‘molecular shock absorber’ action. Certain proteins subject to unfolding force could thus dissipate energy in this fashion to protect from mechanical damage. 1.4.4.3 on-Equilibrium Implications for Free Energy Calculation The non-equilibrium conditions under which proteins are unfolded using the AFM has further implications, both for the unfolding force as well as the potential calculation of free energy from unfolding curves. Within this non-equilibrium transition, energy is lost in the conversion between unfolding a molecule and allowing it to refold. Thus, Gibb’s Free energy cannot be directly calculated for the non-equilibrium unfolding of a protein molecule. Methods exist to calculate the free energy of a reaction occurring far from equilibrium. The Jarzynski 38  equality is one such method; in it, the free energy differences between two equilibrium states is related to the average work done over the irreversible trajectories going from one state to the other.82 This relationship has been utilized in the prediction of free energy surface for the unfolding of RNA83 and the titin I27 domains,84 though controversy exists over the accuracy and applicability of this method within non-equilibrium protein studies.29, 82 1.4.4.4 The Dependence of Unfolding Force on Pull Speed and the Calculation of Unfolding Kinetics Another consequence of the non-equilibrium conditions of single molecule AFM protein unfolding is that the observed unfolding force become dependent on the rate at which the protein is pulled. This dependence in force ramp experiments is expressed as r = dF / dt , where r is the loading rate. Because of this dependence, as well as the high pull rate of most AFM studies, it becomes difficult to extrapolate the unfolding forces observed in vitro to the mechanical stability of proteins within their physiological context, especially considering the non-linear behavior of unfolding force over a large range of pulling speeds. The actual manner in which a protein unfolds may also be affected by the rate at which it unfolds, again making it difficult to extrapolate behavior observed under AFM to that which occurs in vivo. The response of unfolding force to pull rate does make it possible to calculate values of ku and ∆xu by quantifying how unfolding force changes over a range of unfolding forces. This could be done by finding the probability density for unfolding in terms of the spontaneous unfolding rate constant. k T Fu =  B  ∆xu    r∆xu  ln  0   ku k BT      (1.8)  This seemingly straightforward process is complicated by a consequence of the manner in which proteins unfold. Though the pulling speed at which an experiment is done may be kept constant, the actual loading rate acting upon the molecule varies due to the reliance of the applied force on the stiffness of the unfolding molecule. As domains within a polyprotein unfold, this stiffness decreases. Thus, Equation 1.8 may not be directly applied. Instead, unfolding kinetic  39  information is obtained by fitting the changing unfolding force using a Monte Carlo simulation.24 Within this, the unfolding protein is treated as an entropic spring in the same manner as the Worm-like Chain model. Unfolding is modeled as a stochastic, thermally driven process, as described by Eyring rate theory with a two state model determined by a force dependent rate constants. The probability P of observing the unfolding of any part of the polyprotein is described as  P = ku ∆t Where  (1.9)  is the number of individually folded domains in the protein, ku is the unfolding rate  constant, and ∆t is the polling interval utilized during the simulation. The experimental data is fit by running the simulation using altering combinations of kuo and ∆xu until the unfolding force obtain reflects that found experimentally. Fitting in this manner does introduce a degree of ambiguity in that a number of different combinations of ∆xu and kuo may result in the same fit. Differing models, such as one that compares mutant to wild type data,21 have been suggested but are more time intensive; fitting using a Monte Carlo Simulation thus remains the main method by which unfolding kinetic parameters are calculated. 1.4.4.5 Fitting Unfolding Force Distribution Using Unfolding Kinetic Parameters In addition to fitting the speed dependence of unfolding proteins, this Monte Carlo criterion is also utilized to fit the experimental probability distribution of unfolding forces. As previously stated, this distribution is typically broad and skewed towards lower force events demonstrating the effects that thermal motion has upon the unfolding force of the protein. The size and shape of this distribution may be predicted by simulation utilizing the same unfolding kinetic parameters that fit the speed dependence behavior of the molecule. The width of the distribution is directly determined by the unfolding distance ∆xu, where a narrow distribution will result in a larger unfolding distance and vice versa. The unfolding distance ∆xu and the unfolding rate at zero force αo also exhibits a reciprocal effect on the observed unfolding force in that a high unfolding force will result in smaller values of ∆xu and αo. Typically, values for the unfolding rate and distance are resolved from a variety of pulling speed prior to being used to fit a probability distribution at one particular unfolding force. This lessens the effects of 40  experimental broadening effects, such as the presence of spacer elements within the polyprotein structure and cantilever drift. 1.4.4.6 Mechanical Stability is a Kinetic Rather Than Thermodynamic Parameter The treatment of the folded-unfolded transition to this point models the unfolding force, and thus the mechanical stability of the molecule, as a kinetic rather than thermodynamic property. This mechanical stability is determined by the size of the activation barrier ∆GT- as well as the distance between the native and transition state ∆xu. The mechanical stability is obviously kinetic, rather than thermodynamic, in nature. Kinetic properties are dependent on the path taken as a reaction progresses, while thermodynamic properties depend only on the initial and final stages of the reaction. Thus, the thermodynamic stability of the protein is reflected in ∆GU- . This in turn is dependent only upon the state variable describing the system, which is the end-to-end extension of the molecule in the case of protein unfolding. This is clearly not the case, as the unfolding force demonstrates an unambiguous dependence on the rate at which it is pulled. Some proteins demonstrate similarities between their mechanical unfolding rate constants and chemical unfolding parameters, but for most proteins this is not the case. 1.5 The Molecular Determinates of Mechanical Stability as Determined Using Atomic Force Spectroscopy 1.5.1 Emerging Trends in Structural Determinates of Mechanical Behavior Exposed using Single Molecule AFM Although the variety of proteins whose mechanical properties have been investigated using single molecule AFM spectroscopy is small, a number of structural trends in mechanical stability have begun to emerge. High mechanical stability is seen within proteins that exhibit a number of structural features: mechanical stability is often due to a ‘mechanical clamp’ formed by a number of highly localized mechanical hydrogen bonds.85 High mechanical resistance can also be encoded within the hydrophobic core of the protein,86 as well as if the protein is composed of β-strands rather than α-helicies.17 The least mechanically stable proteins tend to be unstructured and β-spiral proteins (such as elastin), while relatively low mechanical stability is also observed within α-helical proteins (such as T4 lysozyme and calmodulin), and an increased 41  degree of stability noted within α-helical bundles (such as myosin II tail87 and spectrin24, 88, 89) and solenoid structures (such as ankyrin B).64 Figure 1.8 Structural Trends in Mechanical Stability Increasing Mechanical Stability  Barnase  Ubiquitin  Calmodulin  T4 Lysozyme 3FnIII  (tenascin)  Figure 1.8 Emerging correspondences between structural elements and mechanical stability. Figure 1.8 illustrates a small section of proteins whose mechanical stability has been previously elucidated utilizing single molecule AFM. Such studies has resulted in the emergence of certain trends between structural composition and mechanical stability, in that proteins with high β-sheet composition tend to demonstrate a higher degree of mechanical stability than do proteins who are composed primarily of α-helices. Mechanical stability can often be ascribed to a small number of hydrogen bonds that act as a ‘mechanical 85 clamp’, resisting mechanical stress.  Mechanical stability cannot simply be ascribed to protein structural composition without consideration of the topology across which mechanical force is applied. This structural topology dictates the mechanism by which mechanical force causes a protein to unfold, and has a large effect on the exhibited mechanical stability of a protein. This effect is exemplified within βstranded proteins. Structures that exhibit a shear mechanical topology, where force is applied orthogonally to hydrogen bonds, are more mechanically stable than ‘zippered’ β-stranded proteins, where force is applied parallel to hydrogen bonds within the structure.29, 90 The relative stability of a ‘shearing’ mechanism as compared to a ‘zippering’ unfolding mechanism is also apparent within the stability disparity observed between proteins with proteins with parallel opposing N and C termini as opposed to proteins with their N and C termini aligned. The heightened mechanical stability of proteins with parallel opposing termini may also be ascribed 42  to the ‘shearing’ mechanism of unfolding, whereby mechanical force is applied across the hydrogen bonds and hydrophobic interactions occurring between force bearing strands. The ‘zippering’ mechanism that occurs within aligned termini structure results in the application of force to sequential bonds as force bearing strands are peeled away. Topological effects delineate the localized nature of mechanical stability, which appears to be dependent on a small number of localized structural interactions rather than on global structural properties. It is not surprising that mechanical proteins that routinely experience physiological mechanical force often exhibit structural and topographical features that impart high mechanical stability. An excellent example of this is Ig domains from the muscle protein titin, whose high resistance to mechanical force stems from terminal A’ and G β-strands arranged into a shear topography. However, proteins that are non-mechanical in nature cannot be grouped as definably in terms of their mechanical stability and corresponding structural attributes as those that are mechanical. The low mechanical stability of barnase and T4 lysozyme, investigated using single molecule AFM, is not surprising considering the similar topologies of these proteins as well as the fact that the ability to withstand mechanical stress is not critical to intrinsic functionality in the same manner as mechanical proteins such as I27. Low mechanical stability is not intrinsic to non-mechanical classes of proteins, as certain non-mechanical proteins have been shown using AFM to exhibit a higher degree of stability. This includes the AFM study of non-mechanical proteins such as green fluorescent protein (GFP), the B1 IgG binding domain of protein L and the B1 IgG binding domain of protein G. All three of these proteins demonstrate high values of unfolding forces, comparable to that of mechanical proteins such as I27. Thus, the non-mechanical nature of a protein does not mean that a protein will demonstrate a low degree of mechanical stability in the same manner that high mechanical stability is necessary for proteins that are mechanical in nature. Investigating the nanomechanical properties of both classes of proteins using single molecule AFM has allowed for the explication of structural and topographical effects on mechanical stability as previously discussed. How mechanical stability is imprinted within the molecular structure of proteins remains unclear, and is aptly demonstrated by the present bottleneck in attempting to rationally increase the mechanical stability of a known protein fold. Though there are many ways in which to decrease this stability, attempting to increase the mechanical stability of a certain protein is difficult. Though use of the AFM has  43  proved an invaluable tool in elucidating molecular trends towards observed mechanical stability, how mechanical stability is molecularly inscribed is still poorly understood.  44  Table 1.1 Mechanical Properties Previously Elucidated using Single Molecule AFM  Protein  SCOP Fold  SCOP Class  Calmodulin  EF hand-like  All α  Barstar  Barstar-like  α+β  T4 Lysozyme  Lysozyme-like  α+β  Barnase  Microbial Ribonuclease  α+β  Green Fluorescent Protein  GFP-Like  α+β  Protein L  β-grasp  α+β  GB1  β-grasp  α+β  Top7 Spectrin (R16) 10 FNIII (fibronectin) 13 FNIII (fibronectin) 12 FNIII (fibronectin) 3 FNIII (tenascin)  Top7 Spectrin repeat-like  Immunoglobulin-like β sandwich Immunoglobulin-like β sandwich Immunoglobulin-like β sandwich Immunoglobulin-like β sandwich Immunoglobulin-like I27 β sandwich Ubiquitin β-grasp 1 FNIII Immunoglobulin-like (fibronectin) β sandwich  Function Nonmechanical Nonmechanical Nonmechanical Nonmechanical Nonmechanical  Unfolding Parallel Force (pN) terminal Reference (Pull speed strands in nm/s) No  <15 (600)  75  Yes  <50 (400)  91  No  50 (400)  92  No  70 (300)  21  No  104 (3000)  24  Yes  136 (400)  29  Yes  180 (400)  18, 93  α+β  Nonmechanical Nonmechanical Novel fold  Yes  155 (400)  85  All α  Mechanical  No  ~30 (600)  94  All β  Mechanical  Yes  75 (400)  86  All β  Mechanical  Yes  89 (400)  17  All β  Mechanical  Yes  125 (400)  17  All β  Mechanical  Yes  120-130 (400)  95  All β  Mechanical  Yes  180 (400)  8  α+β  Mechanical  Yes  203 (400)  90  All β  Mechanical  Yes  220 (600)  17  Table 1.1 Select proteins whose nanomechanical stability has been previously elucidated using single molecule AFM. Proteins previously investigated include a range of both non-mechanical and mechanical proteins of varying size and structural topography. Top7 is a unique case; its fold is novel, designed computationally to be distinct from natural folds. Fold classification according to SCOP criteria.  1.5.2 Investigating The anomechanical Properties of a Mechanical and on-Mechanical Proteins using Single Molecule AFM The following study focuses on observing the nanomechanical properties of both a mechanical protein (Tenascin-X) as well as the non-mechanical enzyme bacillus circulans xylanase (BCX) using the single molecule AFM techniques detailed previously. These proteins 45  demonstrate drastically differing physiological function and sites of expression, and show dramatically dissimilar structures tuned towards contrary functionality. Tenascin-X exhibits the same multimeric design as other mechanical proteins continually exposed to physiological force.96 Its heterogeneous structural design presents a unique model system for disconcerting how these structurally homologous domains contribute to the observed dynamics of the entire heteropolyprotein. Bacillus Circulans Xylanase (BCX) is a member of the endo-beta-(1,4)xylanases family. Unlike the multimeric design of tenascin-X, this enzyme folds into a single domain composed of three β-sheets and one α-helix.97 BCX is a digestive bacterial enzyme critical for the cleavage of long chain xylans from plant cell walls. Unlike Tenascin-X, BCX is functionally non-mechanical nature, demonstrating little obvious structural homology with the mechanical protein tenascin-X. Investigating the nanomechanical properties of each provides a comparative model focusing on how subtle differences in folded structure result in drastically different functionality at the nanoscale, as well as providing general information concerning the folding and unfolding behavior of two dramatically diverse protein folds.  46  1.6 References 1. Bieri, O.; Wildegger, G.; Bachmann, A.; Wagner, C.; Kiefhaber, T., A salt-induced kinetic intermediate is on a new parallel pathway of lysozyme folding. Biochemistry 1999, 38, (38), 12460-70. 2. Pande, V. S.; Rokhsar, D. S., Is the molten globule a third phase of proteins? Proc atl Acad Sci U S A 1998, 95, (4), 1490-4. 3. Pande, V. S.; Grosberg, A.; Tanaka, T.; Rokhsar, D. S., Pathways for protein folding: is a new view needed? Curr Opin Struct Biol 1998, 8, (1), 68-79. 4. Thirumalai, D. W., S. A. (1996) Acc. Chem. Res. 29, 433–439., 1996. 5. Dill, K. A.; Chan, H. 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N., Binding Strength between Cell-Adhesion Proteoglycans Measured by Atomic-Force Microscopy. Science 1995, 267, (5201), 1173-1175. 55. Chilkoti, A.; Boland, T.; Ratner, B. D.; Stayton, P. S., The relationship between ligandbinding thermodynamics and protein-ligand interaction forces measured by atomic force microscopy. Biophys J 1995, 69, (5), 2125-30. 56. Sen, S.; Subramanian, S.; Discher, D. E., Indentation and adhesive probing of a cell membrane with AFM: Theoretical model and experiments. Biophysical Journal 2005, 89, (5), 3203-3213.  49  57. Radmacher, M.; Fritz, M.; Kacher, C. M.; Cleveland, J. P.; Hansma, P. K., Measuring the viscoelastic properties of human platelets with the atomic force microscope. Biophysical Journal 1996, 70, (1), 556-567. 58. Mahaffy, R. E.; Park, S.; Gerde, E.; Kas, J.; Shih, C. K., Quantitative analysis of the viscoelastic properties of thin regions of fibroblasts using atomic force microscopy. Biophysical Journal 2004, 86, (3), 1777-1793. 59. Rief, M.; Oesterhelt, F.; Heymann, B.; Gaub, H. E., Single Molecule Force Spectroscopy on Polysaccharides by Atomic Force Microscopy. Science 1997, 275, (5304), 1295-7. 60. Muller, D. J.; Kessler, M.; Oesterhelt, F.; Moller, C.; Oesterhelt, D.; Gaub, H., Stability of bacteriorhodopsin alpha-helices and loops analyzed by single-molecule force spectroscopy. Biophys J 2002, 83, (6), 3578-88. 61. Desmeules, P.; Grandbois, M.; Bondarenko, V. A.; Yamazaki, A.; Salesse, C., Measurement of membrane binding between recoverin, a calcium-myristoyl switch protein, and lipid bilayers by AFM-based force spectroscopy. Biophysical Journal 2002, 82, (6), 3343-3350. 62. Hinterdorfer, P.; Dufrene, Y. F., Detection and localization of single molecular recognition events using atomic force microscopy. ature Methods 2006, 3, (5), 347-355. 63. Chen, A.; Moy, V. T., Cross-linking of cell surface receptors enhances cooperativity of molecular adhesion. Biophysical Journal 2000, 78, (6), 2814-2820. 64. 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Fersht, A., Structure and mechanism in protein science : a guide to enzyme catalysis and protein folding. W.H. Freeman: New York, 1999; p xxi, 631 p. 79. Bell, G. I., Models for Specific Adhesion of Cells to Cells. Science 1978, 200, (4342), 618-627. 80. Evans, E.; Ritchie, K., Dynamic strength of molecular adhesion bonds. Biophysical Journal 1997, 72, (4), 1541-1555. 81. Evans, E.; Ritchie, K., Strength of a weak bond connecting flexible polymer chains. Biophysical Journal 1999, 76, (5), 2439-2447. 82. Collin, D.; Ritort, F.; Jarzynski, C.; Smith, S. B.; Tinoco, I., Jr.; Bustamante, C., Verification of the Crooks fluctuation theorem and recovery of RNA folding free energies. ature 2005, 437, (7056), 231-4. 83. Liphardt, J.; Dumont, S.; Smith, S. B.; Tinoco, I.; Bustamante, C., Equilibrium information from nonequilibrium measurements in an experimental test of Jarzynski's equality. Science 2002, 296, (5574), 1832-1835. 84. Harris, N. C.; Song, Y.; Kiang, C. H., Experimental free energy surface reconstruction from single-molecule force spectroscopy using Jarzynski's equality. Physical Review Letters 2007, 99, (6), -. 85. Sharma, D.; Perisic, O.; Peng, Q.; Cao, Y.; Lam, C.; Lu, H.; Li, H., Single-molecule force spectroscopy reveals a mechanically stable protein fold and the rational tuning of its mechanical stability. Proc atl Acad Sci U S A 2007, 104, (22), 9278-83. 86. Li, L.; Huang, H. H.; Badilla, C. L.; Fernandez, J. M., Mechanical unfolding intermediates observed by single-molecule force spectroscopy in a fibronectin type III module. J Mol Biol 2005, 345, (4), 817-26. 87. Schwaiger, I.; Sattler, C.; Hostetter, D. R.; Rief, M., The myosin coiled-coil is a truly elastic protein structure. at Mater 2002, 1, (4), 232-5. 88. Law, R.; Carl, P.; Harper, S.; Dalhaimer, P.; Speicher, D. W.; Discher, D. E., Cooperativity in forced unfolding of tandem spectrin repeats. Biophys J 2003, 84, (1), 533-44. 89. Batey, S.; Randles, L. 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P.; Rounsevell, R. W.; Steward, A.; Geierhaas, C. D.; Williams, P. M.; Paci, E.; Clarke, J., Mechanical unfolding of TNfn3: the unfolding pathway of a fnIII domain probed by protein engineering, AFM and MD simulation. J Mol Biol 2005, 350, (4), 776-89. 96. Bristow, J.; Tee, M. K.; Gitelman, S. E.; Mellon, S. H.; Miller, W. L., Tenascin-X - a Novel Extracellular-Matrix Protein Encoded by the Human Xb Gene Overlapping P450c21b. Journal of Cell Biology 1993, 122, (1), 265-278. 97. Plesniak, L. A.; Wakarchuk, W. W.; McIntosh, L. P., Secondary structure and NMR assignments of Bacillus circulans xylanase. Protein Science 1996, 5, (6), 1118-1135.  52  Chapter 2 The anomechanical Properties of Tenascin-X 2.1 Introduction to the Tenascins: A Family of Ubiquitous Extracellular Membrane Proteins 2.1.1 The Extracellular Matrix: A Complex Matrix with Probable Force Transduction Functionality The extracellular matrix (ECM) is the structural entity that lies between cells, representing a complex matrix of fibrous proteins and polysaccharides elements that impart shape, resiliency, and functionality to the tissue it forms. The ECM is critical to a number of processes such as intercellular communication, tissue segregation and organization, sequestration of cellular growth factors,1 and the regulation of processes such as cellular growth, wound healing and fibrosis. The actual composition of the ECM varies dependent upon the organism in which it originates; for example, the main component of the ECM within vertebrates is collagen, while in arthropods and fungi the ECM is primarily composed of chitin. Within vertebrates, the ECM is composed of a complex mixture of proteins and polysaccharides arranged into the basal lamina and interstitial matrix. The basal lamina, or basement membrane, is attached to epithelial cells and is constituted mainly by collagen IV, as well as heparin sulfate proteoglycans, entactin and laminin. The interstitial matrix forms connective tissue; components are secreted via exocytose by resident cells form an interlocking mesh of fibrous proteins and glycosaminoglycans. The predominant structural protein within this mixture is collagen and elastin, interspersed with specialized proteins such as fibronectin and fibrillin as well as proteoglycans such as heparin, chondroitin and keratin sulfate proteoglycans. The incredible structural complexity present within the ECM propend the range of functionality that it exhibits as well as its role as a force transduction medium. The fact that the ECM is continually exposed to force is represented within the structural adaptations exhibited by its constituent proteins. One such protein is fibronectin (Fn) who, with its modular structure, exhibits a large amount of cellular binding sites that provide a diverse array of scaffolding and cell recognition functionality.2 Fibronectin’s role as a force transduction element has been postulated in that the 53 A version of this chapter has been published. Jollymore, A., Lethias, C., Peng, Q., Cao, Y., Li, H. (2009) Nanomechanical Properties of Tenascin-X Revealed by Single-Molecule Force Spectroscopy. J. Mol. Biol. 385, 1277-1286  partial unfolding of the protein could expose cryptic binding sites previously hidden within the folded structure of a domain.3 Thus, it is likely that the ECM is made not only to withstand force, but also to act as a force transduction element to convert force into a relevant biochemical signal. 2.1.2 The Tenascin Family and Their Importance in ECM Organization Through Cell Adhesion The tenascins are a family of cell adhesion proteins that are restrictively expressed within specific sites of the ECM. There are five currently known independent tenascin forms ranging in molecular weight from 230-400 kDa: tenascin C, X, Y, R and W.4 All share a similar structural pattern, beginning with a monomeric subunit composed of an N-terminal hydrophobic ‘head group’ that facilitates polymerization, tandem epidermal growth factor (EGF)-like repeats, tandem fibronectin type III (FnIII) domains and a fibrinogen-like domain at the C terminus.5 Different forms of tenascin exhibit varying numbers of FnIII and EGF domains within their structure. These monomeric forms of tenascin are typically assembled into oligomers via heptad repeats within the C-terminal head unit, and are further stabilized by disulfide bridges in the tenascin-assembly domain.6 As in fibronectin, the FnIII domains within the structure of tenascin contain binding sites for a number of ECM molecules, and a number of ECM proteins, glycosaminoglycans and proteoglycans have been found to bind tenascin with high affinity.7 This binding affinity allows tenascins to act as adhesive proteins, knitting together structural elements of the ECM imparting mechanical strength and elasticity to these tissues. Assigning exact roles to specific forms of tenascin has been difficult, with the effort to assign overarching functionality to the entire family made problematical by seemingly contradictory behavior. For example, tenascin-C and tenascin-X share a similar structure, yet exhibit reciprocal sites of expression and differing specific functionality.8, 9 However, the binding affinity exhibited by the tenascin family imparts organizational properties intrinsic to the functionality of the ECM in a manner that involves constant exposure to mechanical force. How isoforms of tenascin differ in their response to mechanical force, as well as how this force is transduced into a biological signal, could further elucidate the specific mechanotransduction role within the ECM.  54  2.1.3 Tenascin-X is An Integral Connective Protein Within the ECM Tenascin-X (Tn-X) is widely expressed within a variety of mammalian tissues, acting as a connective protein integral to ECM function. It is most widely expressed in fetal skeletal, heart muscle, dermis, testes, nerves and digestive tissue,8-11 with adult expression maintained within skin, joint, heart and blood tissues.12 Interestingly, tenascin-X is more widely expressed within tissues than tenascin-C or tenascin-R,13 pointing to differing functionality among the various tenascin forms. Pinpointing the exact role Tn-X plays within the ECM has been a challenging problem to answer, as has been the case with many of the tenascin species.14 However, the fact that tenascin-X plays an essential role in regulating the structure and mechanical properties of connective tissues is evident from the apparently high binding affinity Tn-X demonstrates for a variety of ECM molecules. This includes collagen fibrils, decorin,15 glycosaminoglycans,16 as well as cell membrane proteins.12 Conclusive evidence that tenascin-X is obligatory for normal functionality of connective tissue comes from a recessive form of the connective disorder EhlersDanlos syndrome caused by a deficiency of or point mutation within Tn-X.17, 18 The EhlersDanlos syndrome (EDS) is characterized by symptoms of hypermobile joints, hypermobile skin, easy bruising and poor wound healing and can generally be attributed to defects in fibrillary collagen metabolism.19 The tenascin-X gene is the first gene outside of the collagen family to be associated with EDS, demonstrating the critical role tenascin-X plays in regulating the mechanical function of the ECM. 2.1.4 Tenascin-X Structure Tenascin-X shares a similar modular structure with other members of the tenascin family. The tenascin-X monomer is formed by an N-terminal head domain, EGF repeats, an FnIII domain region and a carboxyl terminal fibrinogen-like domain.20 The number of EGF and FnIII repeats within the Tn-X structure depends on the species in which it is isolated; in the interest of the following investigation, the focus will be upon bovine tenascin-X. This form of Tn-X is composed of an amino-terminal domain that contains cysteine residues and heptads of hydrophobic amino acids, 18.5 EGF repeats, 30 FnIII domains, and ends with a carboxyl terminal fibrinogen-like repeat.20 As a whole, Tn-X monomers appear to be more flexible than other species of tenascins, possibly owing to the increased number and length of its FnIII 55  domains. Polymerization of monomeric tenascin-X into dimer or trimeric forms is facilitated through N-terminus heptad repeats. This is different than other forms of tenascins, including tenascin-C, which preferentially assembles into hexamers.21-23 Figure 2.1 Bovine Tenascin-X Structure Cysteine-rich domain with heptads repeats  EGF-Like Repeats  FNIII-Like Domains  Fibrinogen like Domain  Figure 2.1 The modular structure of bovine tenascin-X. The bovine tenascin-X monomer is composed of an N-terminal cysteine rich domain with heptad repeats, 18.5 EGF repeats, 30 FnIII domains, and a Cterminal fibrinogen-like domain. Polymerization of monomers into dimers or trimers is facilitated through N-terminal repeats. Monomers of different forms of tenascin differ in the composition of their modular structure as well as the number of monomers that accumulate into their final polymeric form.  2.1.5 Tenascin-X as a Force Spectroscopy Model System Tenascin-X, with its intrinsic mechanical functionality within the ECM, is a good candidate for examination using the single molecule force microscope for a number of reasons. Connective tissues formed by the ECM are continually subject to mechanical in vivo force, where ECM proteins such as Tn-X have evolved to not only withstand such force but also to utilize them by transduction of force into a relevant biological signal. Use of force spectroscopy thus presents an opportunity to investigate nanomechanical properties using a similar set of conditions present physiologically. These advantages were previously used in the explication of the nanomechanical properties of tenascin-C, in which the elastic nature of the molecule, its mechanical stability and kinetics were all elucidated using single molecule AFM.24 Examining these same properties within member of the tenascin family, proteins that follow a similar structural motif but show different functionality and sites of expression, could pave the way to investigating how intrinsic mechanical properties influence exhibited functionality. As well, the FnIII domains present within the structure of Tn-X presents a unique model system in which to comparatively investigate the molecular determinates of folded structure and mechanical stability. The FnIII motif is a ubiquitous one within mammalian proteins, occurring within 2% of all mammalian proteins.25 These individually folded domains exhibit an anti-parallel β-barrel structure that is highly conserved despite low homology observed in the primary structure of 56  these proteins. This seeming dissonance between folded and primary structure is true for modules observed from two differing types of proteins as well as individual domains within the same protein molecule. How these similarities and differences play out into the nanomechanical properties exhibited by different forms of FnIII domains could also elucidate more on how slight molecular differences could cause large mechanical deviations. To this end, FnIII containing proteins whose nanomechanical properties have been elucidated include Tn-C as well as fibronectin. Continued comparative studies between these dissimilar proteins help to examine how their mechanical properties are finely regulated at a mechanical level in order to account for differing functionalities. In order to quantify the nanomechanical behavior of Tn-X as it unfolds under force, four proteins containing all or portions of bovine Tn-X were either constructed or obtained. The four proteins investigated were full length bovine Tn-X, a construct contain only the 30 FnIII domains from Tn-X as well as two polyprotein constructs containing repeats of either the 10th or 11th FnIII domain. 2.2 The anomechanical Properties of Tenascin-X as Demonstrated using Single Molecule AFM 2.2.1 The Unfolding Behavior of Full Length Tn-X Unfolding of full-length Tn-X results in a force extension ‘saw tooth’ pattern in which the individual force peaks correspond to the unfolding of individual domains within the Tn-X structure. The number of events fluctuates depending upon where along the contour of Tn-X attachment between the glass surface and tip occurs, and was observed to be as high as 17 individual events. These unfolding events exhibit a high degree of uniformity in their spacing, with ~24 nm lying between peaks. The force extension curve of full length Tn-X also exhibits a long featureless spacer of up to 150 nm prior to the first unfolding event corresponding to the existence of spacer elements within full-length tenascin structure.  57  Figure 2.2 The Force-Extension Fingerprint of Full Length Tenascin-X Cysteine-rich domain with heptads repeats  EGF-Like Repeats  FNIII-Like Domains  Fibrinogen like Domain  300 pN  50 nm  Figure 2.2 Force extension traces resulting from the mechanical unfolding of full-length tenascin-X. Unfolding of elements within the heterogeneous structure of tenascin-X corresponds to peaks in the sawtooth pattern observed within the force extension trace. Unfolding events were spaced uniformly approximately 24 nm from one another, and were described using a Worm-like chain model (red fit lines). The number of unfolding events varied according to the size of protein randomly picked up by the AFM cantilever tip, with as many as 17 unfolding events appearing within the force extension curve of full length tenascin-X. A long featureless spacer approximately 150 nm in length was noted prior to the first unfolding event corresponding to the presence of inextensible elements within the structure of tenascin-X.  2.2.1.1 Pinpointing the Molecular Origins of Mechanical Features: rT X∆E∆F Because of the heterogeneous modular structure of full length Tn-X, comparing the behavior of specific structural fragments to the overall behavior of the full-length molecule allows for the determination of which structural elements result in observed nanomechanical properties. Comparing the force extension behavior of full length Tn-X to that of its FnIII fragment rTNX∆E∆F reveals that the unfolding peak characteristics of both proteins are nearly identical. This extends to their exhibited ∆Lc and unfolding forces, suggesting that unfolding events occurring within full length Tn-X are due to the mechanical unfolding of FnIII domains. Stretching of rTNX∆E∆F results in characteristic saw tooth-like force-extension curves of as many 17 force peaks (Figure 2.3). Comparing the force extension trace between full length TnX and rTNX∆E∆F shows the longer spacer length in full length Tn-X, a signature that arises as a result of spacer elements within this recombinant protein. These spacer elements contribute to 58  the initial contour length and consist of disulfide bonded EGF domains and non-structured amino acid sequences, elements that are highly disulfide bonded and cannot be unfolded under the few hundred picoNewtons applied using the AFM. The unfolding of the terminal fibrinogen-like domain may also contribute to the initial spacer of the full-length tenascin-X. Figure 2.3 The Force-Extension Characteristics of rTNX∆E∆F FnIII-Like Domains  300 pN  50 nm  Figure 2.3 The unfolding characteristics of the FnIII-domain only Tn-X fragment protein. As in full length Tn-X, the unfolding of fragment FnIII results in force extension curves containing unfolding events spaced ~24 nm apart and of the same approximate magnitude. The mechanical behavior of tenascin-X FnIII domains mirrors that of full length tenascin-X, thus the mechanical properties observed within the full length protein may be ascribed to the FnIII domains present within its structure. Because the FnIIIcontaining fragment rTNX∆E∆F does not contain inextensible elements present within the structure of full-length tenascin-X, a long spacer is not present at low extension as was noted for full-length tenascinX.  2.2.1.2 Determining the Unfolding Force and Contour Length Increment of rT X∆E∆F Compiling the unfolding force from a large pool of single molecule unfolding events results in a force histogram detailing the average unfolding force for Tn-X. Using a number of unfolding events from a number of force extension curves ensures that the average unfolding behavior of Tn-X is shown. Because of the random nature in which molecules are picked up within AFM experiments, different force extension curves of Tn-X will detail the unfolding behavior of different FnIII domains. Correlating unfolding events with the domain demonstrating the unfolding behavior is impossible under nonspecific experimental conditions, thus a compilation of unfolding events report the average behavior among heterogeneous FnIII domains. This average unfolding force was found to be 148 ± 26 pN (n=3100). The 59  heterogeneity between FnIII domains is not reflected in the narrow width of this distribution, suggesting that most of the domains within Tn-X exhibit a similar mechanical stability. This does not rule out the possibility that specific domains may exhibit significantly different values of mechanical stability, as AFM pickup occurs in a random fashion with differing domains demonstrating dissimilar probabilities of being pickup up. Figure 2.4 The Unfolding Force Histogram of Full Length Tenascin-X  500  No. of Events  400 300 200 100 0 0  50  100  150  200  250  300  Force (pN) Figure 2.4 The unfolding force histogram for full-length tenascin-X. The unfolding force histogram details a large number (n=3100) of single molecule unfolding events, and allows for the elucidation of average unfolding force (148 ± 26 pN). The distribution of unfolding forces for tenascin-X is relatively narrow, suggesting that heterogeneous FnIII domains demonstrate similar values of mechanical stability. The -5 -1 distribution of unfolding forces was fit using a Monte Carlo simulation using αo of 9×10 s and ∆xu of 0.34 nm (continuous black line).  Fitting consecutive unfolding domains using the Worm-Like Chain Model allows for the measurement of contour length increment (∆Lc) of individual unfolding domains. As shown in Figure 2.5, the average ∆Lc of all FnIII domains from full length Tn-X demonstrates a narrow distribution with an average value of 27.8 ± 0.7 nm. This is consistent with that found for tenascin-C FnIII domains, which exhibits a ∆Lc of 28.5 ± 4.0 nm24 and agrees with the predicted average of 100 amino acids per FnIII domain. Although the length of individual FnIII domains varies from 88 a.a. to 134 a.a.,20 the narrow distribution of ∆Lc suggests that different FnIII domains are likely to have folded structures that are of similar number of amino acids. Thus, the length variation between different FnIII domains likely reflects discrepancy in linker sequence length between FnIII domains rather than the different number of amino acids present within the 60  folded FnIII structure. This may explain the presence of domains such as the 19th FnIII domain of bovine Tn-X that, at 127 residues, was predicted to be 30 residues longer than other FnIII domains. Because of the narrow distribution in ∆Lc, these thirty residues are likely to act as spacer sequences between the 19th and 20th domains of bovine tenascin-X. Figure 2.5 The Contour Length Increment Histogram of Full Length Tenascin-X  10 0 0  No. of Events  800  600  400  200  0 20  25  30  35  40  ∆Lc (nm) Figure 2.5 The ∆Lc histogram of full length tenascin-X. Despite the heterogeneity present within tenascinX FnIII domains, the distribution of ∆Lc is relatively narrow with an average value of 28.4 ± 1.7 nm, corresponding with the unfolding of ~100 a.a per unfolding event. This suggests that annotated differences in domain size are likely due to linker sequences between similarly sized FnIII modules that do not participate in folding.  That the nanomechanical properties observed within full length Tn-X are due to the unfolding of its FnIII domains may be seen in the correspondence in ∆Lc and unfolding force between full length Tn-X and rTNX∆E∆F. Comparing the average unfolding force for rTNX∆E∆F compiled from a large number of single molecule events as for full length Tn-X again reveals the effects of spacer elements in full length tenascin-X. The average unfolding force of rTNX∆E∆F was found to be 162 ± 26 pN (n=3000, Figure 2.6). A molecule with a longer contour length acts upon the mechanical unfolding force in the same manner as the application of a lower pulling speed; thus it is not surprising that rTNX∆E∆F demonstrates a slightly higher value of average unfolding force. The ∆Lc of rTNX∆E∆F is 27.9 ± 0.6 nm, consistent with that of full length Tn-X (Figure 2.6).  61  Figure 2.6 Mechanical Properties of rTNX∆E∆F 700 300  No. of Events  No. of Events  600 250  200  150  100  500 400 300 200 100  50  0  0 0  50  100  150  200  Force (pN)  250  300  20  25  30  35  40  ∆Lc (nm)  Figure 2.6 The unfolding properties of a fragment protein containing only FnIII domains mirror those of full length tenascin-X. Numerous single molecule unfolding events from the unfolding of the rTNX∆E∆F fragment protein was compiled on the basis of their unfolding force and contour length increment as demonstrated above. The average unfolding force of rTNX∆E∆F was found to be 162 ± 26 pN, while the ∆Lc was determined to be 27.9 ± 0.6 nm. This corresponds closely the unfolding force and ∆Lc as determined for full length tenascin-X, suggesting that FnIII domains within full length structure dictates the mechanical behavior of Tn-X.  2.2.1.3 Speed Dependence of Unfolding Forces Within FnIII Domains The mechanical unfolding kinetics of FnIII domains of full length Tn-X was characterized by carrying out stretching and unfolding experiments at different pulling speeds to measure the speed dependence of mechanical unfolding forces. As shown in Figure 2.7, this dependence is weak: the unfolding force increases from 130 pN at a pulling speed of 50 nm/s to 170 pN at a pulling speed of 5000 nm/s. This weak dependence mirrors that observed for the FnIII domains within Tn-C24,26 but is in sharp contrast with the speed dependence observed in the FnIII domains of fibronectin27 as well as the Ig domains from the giant muscle protein titin.28, 29  A Monte Carlo simulation was used to fit the speed dependence data and to elucidate  unfolding kinetic constants. This simulation was run on the assumption that all the FnIII domains in Tn-X share similar kinetics and unfold in a two state manner in order to estimate the spontaneous unfolding rate constant at zero force (α0) and the distance between the folded state and the mechanical unfolding transition state (∆xu). These parameters were first optimized to fit the speed dependence and further utilized to fit the unfolding force histogram. It was found that a combination of 0.34 nm for ∆xu and 9×10-5s-1 for α0 can describe the speed dependence of 62  unfolding forces and the unfolding force histogram well. Repeating this procedure for rTNX∆E∆F gives values of 0.3 nm for ∆xu and 2 × 10-4 s-1 for α0, contrasting slightly with that observed for FnIII domains within full length tenascin-X. Figure 2.7 Unfolding Kinetics of Full Length and rTNX∆E∆F  A 200  150  100  B  Unfolding Force (pN)  50  0 5  6  7 8 9  100  2  3  4  5  6  7 8 9  7  8  1 000  2  3  4  5  24 0  22 0  20 0  18 0  16 0  14 0  12 0  10 0 2  3  4  5  6  9  2  100 0  P ull S pee d (nm /s )  Figure 2.7 The dependence of unfolding force on pull speed for full length (Figure 2.7 A) and rTNX∆E∆F (Figure 2.7 B) tenascin-X. Both proteins demonstrated a relatively weak correlation between unfolding force and pull speed. This correlation may be utilized to elucidate the unfolding kinetics by fitting experimental data (black squares) using a Monte Carlo simulation (red line). The unfolding behavior of -5 -1 full length tenascin-X may be fit with values of 0.34nm for ∆xu and 9×10 s for α0 (Figure 2.7 A), while -4 -1 the unfolding behavior of rTNX∆E∆F may be described by values of 0.3 nm for ∆xu and 2 × 10 s for α0 (Figure 2.7 B).  2.2.1.4 Folding Kinetics of FnIII Domains In order to measure the refolding kinetics of FnIII domains at the single molecule level, a ‘double pulse’ experiment that repeatedly stretches and relaxes the same molecule may be used.29 This type of experiment is represented in Figure 2.8, demonstrating the unfolding and subsequent relaxation of a single molecule of full length Tn-X. The first ‘pulse’ consists of the forced unfolding of a single molecule trapped between surface and tip. This molecule was fully extended, completely unfolding the FnIII domains within the fragment picked up (trace 1). The 63  number of domains within the extended fragment is equal to the number of peaks within the force extension curve (  total).  After the full extension of the polyprotein was achieved, refolding  was then initiated by relaxing the molecule back to zero extension. The protein was allowed to refold at this resting position for a fixed period of time ∆t. After this time, the molecule was then re-stretched in order to measure its force extension profile. This second force extension curve demonstrates the number of domains that have managed to refold and regain their mechanical stability within the time ∆t by measuring the number of domains unfolding during this second extension (  refold).  If ∆t is lengthened, the probability that more FnIII domains will refold is  higher than at smaller time intervals. Thus, varying this ∆t value and measuring the proportion of domains able to refold during this time allows for the direct measurement of folding kinetics. This was accomplished by plotting the folding probability of full length Tn-X versus the relaxation time interval ∆t. The time course of FnIII folding may be well described using a double exponential model:  refold  (  )  (  = A 1 − e − β1t + (1 − A) 1 − e − β 2 t  )  (2.1)  total  where  refold/ total  is the folding probability of FnIII domains at a relaxation time t, and β1 and β2  refer to the folding rate constants. Thus, full length Tn-X exhibits biphasic folding behavior, with 60% of FnIII domains fold with a rate constant of 22 s-1 and 40% of these domains folded at a rate of 3 s-1. This biphasic behavior may be attributed to the heterogeneous composition of the FnIII domains constituting its structure, where differing domains may exhibit differing folding behavior.  64  Figure 2.8 Folding Kinetics of Full Length Tenascin-X A (1)  B ∆t=0.02 s 1.0  (2)  Folding Probability  (3)  ∆t=0.12 s  ∆t=1 s  0.8  0.6  0.4  200 pN  0.2  50 nm  0.0 0.0  0 .5  1.0  1.5  2.0  ∆t (s)  Figure 2.8 Mechanical folding kinetics of FnIII domains of tenascin-X. (2.8 A) The folding kinetics of FnIII domains can be measured using a double-pulse protocol. In the first pulse, tenascin-X was picked up and stretched to unfold all the FnIII domains within the fragment. The number of unfolding force peaks allows for the determination of the number of domains present within the attached molecule of protein (trace 1). After complete unfolding, the unfolded protein chain was quickly relaxed to zero extension (trace 2) and allowed to refold at zero extension for time ∆t. The protein was then re-stretched; the number of unfolding force peaks in the resultant force–extension curve (trace 3) indicates the number of FnIII domains that managed to refold within relaxation time ∆t. By varying the relaxation time ∆t, the folding kinetics of FnIII domains of tenascin-X can be measured. (Figure 2.8 B) Folding kinetics of FnIII domains. The profile of folding probability of FnIII domains (Nrefold/Ntotal) versus relaxation time (∆t) can be described by a double −1 −1 exponential distribution with a folding rate constant of 22 s for the fast phase and 3 s for the slow phase.  2.2.2 Mechanical Unfolding and Folding Dynamics of Individual FnIII Domains of Tenascin-X Unfolding full length and FnIII fragment tenascin-X results in the elucidation of mechanical unfolding and refolding parameters as demonstrated. However, such information is intrinsically averaged due to the heterogeneity observed between different FnIII domains. Thus, it becomes impossible to concretely assign the specific behavior of a domain from the properties observed from unfolding all FnIII domains. In order to obtain such information, it becomes necessary to investigate the mechanical unfolding and folding dynamics of FnIII domains one at a time.  65  Two consecutive domains, the tenth and eleventh domain of bovine tenascin-X, were chosen as model systems to illustrate some of the general features common among FnIII domains. These investigations centered on the construction and use of two octameric heteropolyproteins containing either the tenth or eleventh domain alternating with the protein GB1 (Figure 2.9). Stretching of both (GB1-TNXfn10)4 and (GB1-TNXfn11)4 polyproteins resulted in characteristic saw tooth-like force extension curves that clearly demonstrating the presence of both proteins. GB1 has been well characterized using single molecule AFM, exhibiting a distinct ∆Lc of ∼18 nm and an unfolding force of ∼180 pN. It thus serves as an obviously definable molecular fingerprint for identifying the mechanical features of the Tn-X domains, where peaks that occur with an ∆Lc of 18 nm are due to the unfolding of GB1. Unfolding events occurring at low extension tend to demonstrate an ∆Lc of ~28nm, corresponding to the unfolding of Tn-X FnIII domains while unfolding events at higher extension tend to show the approximate 18nm ∆Lc indicative of GB1 unfolding.  66  Figure 2.9 Mechanical Unfolding of FnIII10 and FnIII11 Polyproteins  A  GB1  FnIII10  GB1  FnIII10  GB1  FnIII10  GB1  FnIII10  18 nm  28 nm  200 pN 50 nm  B  GB1  FnIII11  28 nm  GB1  FnIII11  GB1  FnIII11  GB1  FnIII11  18 nm  200 pN 50 nm  Figure 2.9 The mechanical unfolding of polyproteins containing the tenth and eleventh tenascin-X FnIII domains. Two octameric polyproteins were constructed by alternating the molecular fingerprint GB1 with either the tenth FnIII domain (Figure 2.9 A) or the eleventh FnIII domain (Figure 2.9 B). Mechanically unfolding either of these polyproteins resulted in a characteristic sawtooth unfolding pattern that revealed the presence of both types of modules within the polyprotein structure. GB1 unfolding events were ascribed to those that exhibited a representative ∆Lc of ~18 nm and unfolding forces of ~180 pN and typically occur at higher force extension (unfolding events in red). Unfolding events with ∆Lc of ~28 nm typically occur at the beginning of force extension curves, and were ascribed to the unfolding of both FnIII10 (Figure 2.9 A, green unfolding events) and FnIII11 (Figure 2.9 B, blue unfolding events).  67  2.2.2.1 The Mechanical Unfolding Properties of T Xfn10 That the unfolding events of the TNXfn10 portion of (GB1-TNXfn10)4 may be clearly identified by its ∆Lc allows for the compilation into an unfolding force distribution as was done for full length Tn-X and rTNX∆E∆F (Figure 2.10). The average unfolding force of TNXfn10 was thus demonstrated to be 170 ± 22 pN, while a similar distribution for GB1 revealed an average unfolding force of ∼180 pN. Immediately apparent is that the average unfolding force for TNXfn10 is higher than the average force of FnIII domains measured in tenascin-X. Unfolding parameters such as α0 and ∆xu were elucidated according to the dependence of mechanical unfolding forces on pulling speed. It was found that the parameters used to describe the average behavior of FnIII domains in Tn-X can also be used to describe the speed dependence of force as well as the probability distribution of unfolding forces of TNXfn10 (Figure 2.10). This is consistent with the conclusion that the majority of FnIII domains demonstrates similar mechanical unfolding kinetics and may be fit using similar values of α0 and  ∆xu. Figure 2.10 Unfolding Force Histogram and Kinetics of TNXfn10 B  A 160  250  Unfolding Force (pN)  No. of Events  140 120 100 80 60 40 20  200  150  100  50  0 0  50  100  150  200  250  Unfolding Force (pN)  300  350  2  100  3  4  5  6  7  8 9  2  3  4  5  1000  Pull Speed (nm/s)  Figure 2.10 The average unfolding force and unfolding kinetics of TNXfn10 elucidated using single molecule AFM. The unfolding force histogram formed by compiling unfolding events of TNXfn10 domains, shown in Figure 2.10 A, demonstrates the average unfolding force of the tenth FnIII domain of tenascin-X to be 170 ± 22 pN. This average unfolding force is apparently higher than that of full length or rTNX∆E∆F tenascin-X. Unfolding kinetics of TNXfn10 was determined by exploiting the dependence of unfolding force on pull speed, as shown in Figure 2.10 B. Fitting experimental data was done using a Monte Carlo simulation with the same values of ∆xu and α0 as that used to fit full length tenascin-X (0.34 -5 -1 nm and 9×10 s respectively).  68  The refolding kinetics of TNXfn10 was also elucidated using the same double pulse protocol as for full length tenascin-X (Figure 2.11). Folding of TNXfn10 may be readily described by a single exponential distribution with a folding rate constant β of 3 s-1. This folding rate mirrors the slower folding rate observed within the averaged behavior of all FnIII domains in tenascin-X. That the mechanical behavior of TNXfn10 so closely mirrors that of averaged FnIII domains demonstrates the congruity between individual domain properties and that of its polyprotein. Figure 2.11 Folding Kinetics of TNXfn10  Folding Probability  1.0 0.8 0.6 0.4 0.2 0.0  0  1  2  3  4  5  ∆t (s)  Figure 2.11 The folding kinetics of TNXfn10. These kinetics were elucidated using the same double pulse protocol as was described previously for full length tenascin-X. Briefly, a single molecule of the (GB1TNXfn10)4 polyprotein was extended such that all domains were unfolded. Counting the number of unfolding events within the resultant force extension curve revealed the number of domains within the extended polyprotein, which was subsequently allowed to relax to zero extension. After a certain amount of time ∆t at zero extension, the protein was re-extended to determine the number of domains that had refolded during this time. Varying the amount of refolding time elucidates the folding probability -1 distribution, which was fit using a single exponential function with folding rate constant β of 3 s .  2.2.2.2 The Mechanical Properties of T Xfn11 As for TNXfn10, the mechanical unfolding of TNXfn11 is clearly identified from force extension curves of (GB1-TNXfn11)4 using its characteristic value of ∆Lc. These extension curves revealed the saw tooth-like behavior of these molecules under force, exhibiting TNXfn11 events at low extension and ∆Lc of 28 nm (Fig 2.9). As was the case for TNXfn10, GB1 occurred at higher extension with an ∆Lc of 18 nm and an average unfolding force of 180 pN. 69  Compiling TNXfn11 unfolding events into an unfolding probability distribution reveals an average unfolding force of 122 ± 17 pN (Fig 2.12). This average unfolding force is obviously lower than that determined for either the averaged FnIII domains or for the individual TNXfn10. Unfolding kinetic parameters were determined according to the dependence of unfolding force on pulling speed (Fig 2.12) and these parameters used to fit the probability force distribution. Unlike the TNXfn10 domain, where mechanical unfolding of the isolated domain mirrored the average behavior in FnIII domains, TNXfn11 revealed differing unfolding kinetics. The speed dependence of TNXfn11 was best fit using a ∆xu of 0.35 nm and αO of 2×10-3 s-1. Figure 2.12 The Unfolding Force and Unfolding Kinetics of TNXfn11  A  B 250  Unfolding Force (pN)  160  No. of Events  140 120 100 80 60 40 20  200 150 100 50 0  0  5  0  50  100  150  Force (pN)  200  250  300  350  6  7 8 9  100  2  3  4  5  6  7 8 9  2  3  1000  Pulling Speed (nm/s)  Figure 2.12 The unfolding force histogram and unfolding kinetics of TNXfn11. Figure 2.12 A details the unfolding force distribution of TNXfn11, where the average unfolding force of TNXfn11 was determined to be 122 ± 17 pN. The unfolding force distribution of TNXfn11 was uniformly lower than any of the other tenascin-X proteins examined. Figure 2.12 B shows the dependence of unfolding force on pulling speed for TNXfn11. This distribution was fit using a Monte Carlo simulation (red line), in order to elucidate -3 -1 values of 0.35 nm of ∆xu and 2×10 s for αO, which was subsequently used to fit the unfolding force distribution (Figure 2.12 A, black continuous line).  This kinetic disparity was also apparent within the folding behavior of TNXfn11, which was elucidated using the same double pulse protocol as was previously described. Folding of TNXfn11 was well described using a single exponential distribution with a folding rate constant  β of 31 s-1. This folding behavior is also not represented within the averaged folding properties of FnIII, representing a faster folding phase than any previously exhibited within any of the tenascin-X models.  70  4  5  Figure 2.13 Folding Kinetics of TNXfn11  Folding Probability  1.0 0.8 0.6 0.4 0.2 0.0 0  1  2  3  4  5  ∆t (s) Figure 2.13 The folding kinetics of TNXfn11 as elucidated using a double pulse refolding protocol as previously described. The folding probability distribution of TNXfn11 may be fit using a single exponential -1 fit with a folding rate constant β of 31 s .  2.3 Discussing Tenascin-X as an Elastic Protein with Well Defined Mechanical Properties That tenascin-X is intrinsically elastic in nature of evident from its force extension trace; consecutive FnIII domains within its structure may be reversibly unfolded under force without apparent loss of mechanical stability within the refolded protein structure. Quantifying mechanical properties such as unfolding and folding kinetic parameters of FnIII domains of tenascin-X allows for the comparison to be made across different forms of tenascins and other FnIII containing proteins. As discussed previously, the five known tenascins constitute a family of highly conserved extracellular matrix proteins. All members of the tenascin family display a similar architecture despite their low sequence homology, with FnIII domains constituting a major portion of tenascin structure. The FnIII domains are prime structural candidates for providing unfolding, and it is not surprising that this is apparent in the unfolding of both tenascin-X demonstrated here as well as for the elastic behavior of tenascin-C examined previously.24, 26 Although the other three forms of tenascin, tenascin –R, tenascin-W and tenascin-Y, remain to be investigated, their basic behavior may be predicated based on properties observed within AFM studies on tenascinC and tenascin-X. Members of the tenascin family are likely to be elastic, with their FnIII 71  domains acting as structural elastic elements mediating their force induced unfolding. It makes sense that the FnIII domains are the main determinates of elastic behavior, as other general structural elements within the tenascins are obviously not likely to be extended via AFM. This includes highly disulfide bonded EGF domains, which remain folded under force applied using AFM and contributes to the initial contour length of tenascins. Thus, if the elastic properties of FnIII domains may be correlated throughout the tenascin family, it is possible that other forms of functionality may also be interrelated. It has been proposed that the reversible unfolding of FnIII domains in tenascin-C may effectively extend the life time of adhesion bonds that this protein mediates.24 Tenascin-X, like other members of the tenascin family, has been demonstrated to adhere to a number of ECM substituents, such as heparin, decorin, and collagen fibrils, and helps to provide desired mechanical strength and elasticity to connective tissues as well as regulating the fine structure of these tissues. Binding to a variety of ECM proteins appears to be a trait shared by members of tenascin family; thus, it is possible that the mechanism proposed for tenascin-C FnIII domains is common to all tenascin FnIII domains including tenascin-X and unfolding of FnIII domains can help to promote tenascin-involved adhesion bonds. Reporting the properties of full length tenascin-X is a step towards comparing its mechanical characteristics; however, in order to obtain detailed information on specific FnIII domains they must be investigated individually. Examining the behavior of isolated domains, such as was done for TNXfn10 and TNXfn11, offers information about how specific molecular determinates easily isolated at the domain level contribute to the observed mechanical stability. These studies pave the way to evaluating the mechano-phenotypic effect of disease causing mutations, such as those found within a few FnIII domains and linked to connective disorders.18, 30  Thus, comparing the mechanical behavior of the two isolated domains reveals some  surprising variations. Both domains are of a similar annotated size, with both domains containing close to 100 amino acids. The mechanical behavior of TNXfn10 most closely resembled the averaged properties observed from the unfolding of all Tn-X domains; both the unfolding and folding characteristics fell into those observed within averaged FnIII domains. The only discrepancy was the observed increase in average unfolding force between that of TNXfn10 and both fragment and full length tenascin-X; this increase may due to a slight amplification in mechanical stability within this domain, but could also be due to the lack of  72  significant spacers within the polyprotein construct. This same effect is observed in the average unfolding force of rTNX∆E∆F compared to full length tenascin-X, which increased the average unfolding force of rTNX∆E∆F by a similar amount as that observed within TNXfn10. The behavior of TNXfn11 was not as clearly reflected within that observed for averaged FnIII. When compared to the other tenascin-X proteins examined, the average unfolding force is significantly lower. This decrease is unexpected, especially considering the spacer effects previously discussed in reference to the increased unfolding force observed within rTNX∆E∆F and TNXfn10 that would presumably also apply in the case of TNXfn11. An observed decrease in apparent mechanical stability could be due to a number of causes. Isolating a specific domain may cause the mechanical stability of the folded structure to decrease if the domain is subject to stabilization effects of another folded domain within close proximity. Stabilization effects from neighboring domains have been previously observed in closely associated but independent domains.31 TNXfn10 and TNXfn11 are closely associated through a heparin binding site that involves sites on both domains, an association that may be critical in fully achieving a properly mechanically stable fold for both. That TNXfn11 also diverges from observes TNX behavior could also be an intrinsic, though anomalous, property of this specific domain. Although the force probability distribution of averaged FnIII domains shows a relatively narrow distribution, this does not preempt the fact that domains unfolding at forces far from the average observed could also be present within the total FnIII structure. Pickup of domains is probabilistic along all 30 of the FnIII domains under investigation, where certain domains may be underrepresented within the distribution. As noted, observing the unfolding behavior of all 30 FnIII domains as for full length tenascin-X and rTNX∆E∆F reports the average behavior of all domains, where anomalous characteristics of exceptional domains may be lost within the average.  73  2.3.1 Comparison of the Mechanical Features of FnIII Domains Between Tenascin-X and Tenascin-C A comparison between the mechanical properties of FnIII domains from tenascin-X and tenascin-C reveals that these proteins exhibit some interesting common as well as distinguishing mechanical features. These FnIII domains originating from either tenascins show a large degree of intra- and inter-protein sequence diversity; however, FnIII domains from both proteins demonstrate relatively congruous average unfolding force and mechanical unfolding kinetics. This is not the case for FnIII domains deriving from fibronectin, which exhibit a marked hierarchy when unfolded using force.27 This difference cannot be ascribed to differing sequence homologies, which is similar between diverse types of FnIII domains. Instead, it is possible that similar mechanical stabilities among FnIII domains are a unique characteristic for the tenascin family. Deciphering the mechanism by which these mechanical features are encoded within the FnIII domains has implications towards the rational tailoring of mechanical stability, and is an aspect of study where steered molecular dynamics (SMD) simulations would be of tremendous help. Preliminary study of the unfolding of fibronectin FnIII domains is a step in this direction,32-34 where comparative study of the mechanical design of both tenascins would provide further information as to how differing molecular features give rise to observed mechanical behavior. Similarly, both tenascin-X and tenascin-C exhibit heterogeneous folding behavior (Figure 2.14) that manifests itself within observed double exponential kinetics. This is in line with the proposed mechanism whereby such heterogeneity is an intentional feature present within tandem modular proteins to prevent misfolding. Despite these similarities, tenascin-C and tenascin-X display some notable differences within their mechanical behavior. The most evident is the apparent divergence in the exhibited mechanical stability of their FnIII domains: FnIII domains from tenascin-X are consistently stronger than those from tenascin-C. Considering the low FnIII sequence homology between tenascin-C and tenascin-X, these results imply that the FnIII domains within the same type of tenascin likely evolved similar mechanical stability distinct from that of other forms of tenascin. Different forms of tenascins play various organizational roles within the ECM, thus divergences within mechanical stability between forms of tenascin may encode for these differing functional roles. The observable mechanical differentiation may thus be specifically engendered in order to  74  impart specific functionality to the dissimilar forms of tenascin in much the same way the mechanical properties of fibronectin differ from the general characteristics of the tenascins. Tenascin-X and tenascin-C also differ in the manner in which they refold after forced unfolding, where tenascin-C reports a significant increase in mis-folding frequency. Previously, investigations of tenascin-C reported that 10% of neighboring FnIII domains was observed to refold into dimeric misfolding states after mechanical unfolding. The appearance of misfolded states is shown by a ‘skip’ in unfolding events within the force extension curve of tenascin-C. This skip results in an ∆Lc of approximately 60nm, slightly larger than twice the ∆Lc of two properly folded FnIII domains.12, 35 The folding of FnIII within tenascin-X appears to be more robust: in more than 500 refolding events, only 2 putative misfolding events were observed. The sharp contrast in the folding fidelity of FnIII domains between tenascin-X and tenascin-C raises the question of how the folding fidelity of such similar tandem modular proteins is encoded within their amino acid sequences. It has been previously postulated that high sequence homology is likely a cause for misfolding and aggregation within tandem modular proteins.36 This does not explain the behavior observed between the two forms of tenascin, who both exhibit similarly low values of FnIII sequence homology. Hence, FnIII domains from tenascin-C and tenascin-X may represent a valuable model system in which to examine the molecular determinants underlying these differing mechanical properties and what the biological significance of this diversity may be.  75  Figure 2.14 The Folding Fidelity of Tenascin-X is Greater than Tenascin-C  A  B  Figure 2.14 FnIII domains of tenascin-X show high fidelity in folding. Figure 2.12 A demonstrates the refolding traces of tenascin-C showing misfolding events. After mechanical unfolding, about 10% FnIII domains of tenascin-C can refold with their neighboring FnIII domains to form a misfolded state, which give rise to ∆skip of ~60 nm, which is slightly bigger than twice the ∆Lc observed for the individual correctly folded FnIII domains. Figure 2.12 B demonstrates typical refolding traces of tenascin-X. After mechanical unfolding, FnIII domains refold to regain their mechanical stability and show ∆Lc that is identical to that of FnIII domains upon their first unfolding.  2.4 Conclusion This study characterizes the elastic behaviors of full length tenascin-X as well as identifies the specific properties of its constituent FnIII domains. These results show that tenascin-X is an elastic protein whose mechanical properties are governed by the reversible unfolding of its FnIII domains. When the folding and unfolding behavior of tenascin-X is compared to the previously elucidated behavior of tenascin-C, a number of similarities as well as a few critical differences in behavior are revealed. That much of the behavior between the two proteins is shared may be a result of a similar structure and functionality present for all tenascin family members, while divergences in behavior impart specific biological functionality onto each 76  particular tenascin form. These results pave the way for further studies to investigate how the mechanical properties of tenascin-X regulates its observed physiological functionality and how the mechanical performance of this protein is affected by disease causing mutations. 2.5 Investigating the Properties of Tn-X: Materials and Methods 2.5.1 Full Length and All FnIII Tn-X Fragment Both the recombinant full length and all FnIII fragment of bovine tenascin-X were obtained in collaboration with Claire Lethias. These proteins were prepared as described previously,37 produced in HEK293 cells and secreted in the culture medium. Purification was done in two chromatographic steps, first a heparin affinity column followed by a Q-Sepharose column. 2.5.2 Construction of FnIII Containing Polyproteins 2.5.2.1 Construction of Genes Containing FnIII10 and FnIII11 Two constructs containing Tn-X FnIII domain fragments were constructed for study using AFM, one containing the tenth FnIII domain and the other containing the eleventh FnIII domain of bovine tenascin-X. These constructs were designed to alternated the protein GB1 with the tenascin domain in a manner previously described.38 Integrating GB1 into the polyprotein serves a number of advantageous functions; firstly, GB1 has been well characterized using AFM and is known to unfold with a ∆Lc of ~18nm. GB1 can thus be used as a fingerprint in the force extension of the GB1 Tn-X chimera, allowing for the clear identification of force extension peaks caused by the unfolding of the FnIII domains. Secondly, GB1 proteins are known to be easily expressed using the E.Coli strains and gene vectors detailed below, ensuring simplicity in procuring the desired polyprotein construct. Both polyproteins were constructed in the same manner, beginning with amplification of the desired domain from a plasmid encoding TNXfn9-11 using standard polymerase chain reaction (PCR). Choice of primers used during the PCR reaction were designed to ensure that the desired domain was correctly amplified in the case of both TNXfn10, with sense primer (CGT  77  GGA TCC CTG CTC TTT GGC ATC CAA GAT) and GGG AGA TCT TAA TAG GGT ACC GAG) AGA GCT GTG GAT GAG GCT GG) and CAG GGT CAC GGC GAT GGC).  antisense primer (CCC TCA AAA CCA CGT CTG  and TNXfn11, with sense primer (CGT GGA TCC CGC  antisense primer (CTC GGT ACC CTA TTA AGA TCT CTG  Using primers that either slightly overlapping neighboring  domains (as in the case of TNXfn10) or all of the existing annotation of the domain (as in TNXfn11) ensures that the entire domain is contained within the gene construct, eliminating the possibility of encountering effects due to incorrect boundary choices. These primers were also designed such that the amplified gene contained 5’ BamHI and 3’ KpnI and BglII restriction enzyme sites. Successful amplification of the desired domain was confirmed using direct DNA sequencing. Both amplified gene fragments containing Tn-X FnIII fragments were subsequently ligated using T4 DNA ligase into a pUC19 DNA vector containing the GB1 gene at its 5’ end, creating a [GB1-TNXfn] dimer. The successful dimerization was verified using a small portion of the gene in solution. This small amount of plasmid in solution was incubated at 37̊C for two hours with the restriction enzymes BamHI and KpnI, which act to cut the dimerized gene out of the pUC19 plasmid. The size of the isolated linear gene fragment was verified by running the gene fragment as well as a standardize gene ladder on a 1% agarose gel stained with ethidium bromide. The size of the construct was easily determined from the known size of both GB1 as well as the Tn-X fragment, and once the size was verified, the remaining plasmids were transformed into XLI Blue competent cells. This transformation stage was utilized to increase the amount of plasmid DNA containing the pertinent gene constructs by exploiting natural replication mechanisms of the cell. Plasmid transformation requires that the plasmid is successfully introduced through the nuclear membrane into the nucleus of the cell, where the machinery necessary to replicate the plasmid DNA is located. This was done via a heat shock stage that acts to open the nuclear pore of the E. Coli host and admit plasmid DNA into the nucleus for replication. In order to increase the concentration of the engineered plasmid as well as to ensure its purity, the gene was created such that a gene transcribing for antibiotic resistance is also engineered within the plasmid in addition to the desired gene. Thus, growing transformed cells in an incubated medium containing both nutrient broth and antibiotics ensures that only cells containing the correct gene survives and replicate. The amplified gene was purified for  78  further use by lysing the cell membrane, centrifuging the sample to separate the soluble plasmid DNA from insoluble cell components such as the cellular membrane, and binding the plasmid DNA to a silica DNA-affinity column. The plasmid DNA was then eluted and stored at a temperature of –80˚C. Construction of a tetrameric gene product began in the same manner as the analysis of the dimer size; a dimeric gene insert was created by incubating the plasmid DNA with BamHI and KpnI restriction enzymes. This formed a dimeric linearized gene product flanked by 5’ and 3’ overhanging ‘sticky ends’ that are the result of restriction enzyme digestion. Incubating a portion of the dimeric plasmid with KPNI and BglII restriction enzymes created a DNA vector containing the Tn-X dimer; this step linearized the dimeric plasmid. Both of the incubation products were run on an 1% agarose gel, a step meant to separate the desired gene product from the enzymes used as well as to differentiate between the desired linearized DNA and the small fraction of super coiled DNA produced as a byproduct of the restriction enzyme process. The desired bands were excised from the gel, solubilized in an aqueous buffer, and purified using a DNA affinity column. The short insert DNA containing the [GB1 TNX] dimer was ligated into the linearized pUC19[GB1 TNX] plasmid using T4 DNA ligase, and the newly formed tetrameric plasmid transformed into XLIBlue competent cells. This process was repeated once more using the tetrameric gene construct to create an octameric chimera. The last step involved extraction of the octameric insert using BamHI and KPNI restriction enzymes and subsequent ligation of this insert into a pQE80L DNA plasmid cut with KPNI and BglII. The transition from pUC19 to pQE80L was done as the pQE80L plasmid contains elements necessary for subsequent protein expression steps, including IPTG induction elements such as a IPTG operator. In order to ensure that the final pQE80L[GB1TNX]4 vector contained the appropriately sized insert, and that the previous rounds of restriction digestion and ligation was successful, a small fraction of the octameric pQE80L[GB1TNX]4 was digested with BamHI and KPNI and run on an agarose gel to verify the insert size.  79  2.5.2.2 Expression of the FnIII Containing Polyprotein The expression of cloned genes containing either FnIII10 or FnIII11 was accomplished by over expressing both within DH5α competent cells. Cells containing the desired plasmid were grown to optical density of approximately 0.6 (at 600 nm) at 37˚C. At this time, 1M of IPTG was added to induce expression of the [GB1 TNX]4 gene and the culture allowed to grow for an additional four hours. After this induction period, cell medium was removed from heat and cell s lysed using 1M lysozyme in solution. Addition of DNase and RNase solutions summarily degraded DNA and RNA from the lysis solution. The lysed protein solution was then centrifuged at approximately 4000 rpm. If induction and expression of the protein was successful, then the soluble protein of interest will be within the soluble supernatant. Components of the pellet (inclusion body) include components of the cell membrane and other insoluble components of the lysed cell. Purification of the pertinent protein product from the centrifuged supernatant was accomplished by applying the filtered supernatant to a column containing a Talon Co2+ Resin. Purification of the desired protein product was accomplished via binding between a poly-histidine tag expressed on the N-terminus of the desired protein construct and the Talon Affinity Co2+ Resin column (Clontech). Elution from the column was accomplished using a phosphate buffered saline (PBS) solution containing imidazole, which causes the release of protein from the resin due to the increased affinity between the more concentrated imidazole solution and the metal affinity column. The presence of the protein construct within the elution fraction was ascertained by running an aliquot of solution on a SDSPAGE gel. The position of the resultant protein band within the SDS-PAGE gel was compared to that of a prestained protein marker to determine the size of the protein fragment within solution. The concentration of protein was then determined via UV absorption, with the TNXfn10-containing protein exhibiting a concentration of 0.3 µg/mL and the TNXfn11containing protein at a concentration of 1.0 µg/mL. 2.5.3 Single Molecule AFM Experiments Single-molecule AFM experiments were carried out on a custom-built AFM, which was constructed as described previously.38 Each individual AFM cantilever was calibrated using the equipartition theorem before and after each AFM experiment; typical spring constant values for 80  the cantilever ranged from 50-80 pN/nm. Approximately 1.0 µl of protein solution in PBS was deposited onto a clean glass coverslip covered by phosphate-buffered saline buffer (∼50 µl) and allowed to adsorb for approximately 5 min. During AFM experiments, the AFM tip was first brought into contact with the glass coverslip with a contact force of 5 nN. Upon withdrawing the AFM tip from the glass surface, tenascin fragments attach to the cantilever tip through nonspecific interactions; these interactions were strong enough to allow for subsequent experimental rounds of stretching and relaxation.  81  2.6 References 1. Kumar, V.; Abbas, A. K.; Fausto, N.; Robbins, S. L.; Cotran, R. S., Robbins and Cotran pathologic basis of disease. 7th ed.; Elsevier Saunders: Philadelphia, 2005; p xv, 1525 p. 2. Smith, M. L.; Gourdon, D.; Little, W. C.; Kubow, K. E.; Eguiluz, R. A.; Luna-Morris, S.; Vogel, V., Force-induced unfolding of fibronectin in the extracellular matrix of living cells. PLoS Biol 2007, 5, (10), e268. 3. Vogel, V., Mechanotransduction involving multimodular proteins: converting force into biochemical signals. Annu Rev Biophys Biomol Struct 2006, 35, 459-88. 4. Tucker, R. P.; Drabikowski, K.; Hess, J. F.; Ferralli, J.; Chiquet-Ehrismann, R.; Adams, J. C., Phylogenetic analysis of the tenascin gene family: evidence of origin early in the chordate lineage. BMC Evol Biol 2006, 6, 60. 5. Gulcher, J. R.; Nies, D. E.; Alexakos, M. J.; Ravikant, N. A.; Sturgill, M. E.; Marton, L. S.; Stefansson, K., Structure of the human hexabrachion (tenascin) gene. Proc atl Acad Sci U S A 1991, 88, (21), 9438-42. 6. Kammerer, R. A.; Schulthess, T.; Landwehr, R.; Lustig, A.; Fischer, D.; Engel, J., Tenascin-C hexabrachion assembly is a sequential two-step process initiated by coiled-coil alpha-helices. J Biol Chem 1998, 273, (17), 10602-8. 7. Jones, F. S.; Jones, P. L., The tenascin family of ECM glycoproteins: structure, function, and regulation during embryonic development and tissue remodeling. Dev Dyn 2000, 218, (2), 235-59. 8. Geffrotin, C.; Garrido, J. J.; Tremet, L.; Vaiman, M., Distinct tissue distribution in pigs of tenascin-X and tenascin-C transcripts. Eur J Biochem 1995, 231, (1), 83-92. 9. Matsumoto, K.; Saga, Y.; Ikemura, T.; Sakakura, T.; Chiquet-Ehrismann, R., The distribution of tenascin-X is distinct and often reciprocal to that of tenascin-C. J Cell Biol 1994, 125, (2), 483-93. 10. Bristow, J.; Tee, M. K.; Gitelman, S. E.; Mellon, S. H.; Miller, W. L., Tenascin-X: a novel extracellular matrix protein encoded by the human XB gene overlapping P450c21B. J Cell Biol 1993, 122, (1), 265-78. 11. Burch, G. H.; Bedolli, M. A.; McDonough, S.; Rosenthal, S. M.; Bristow, J., Embryonic expression of tenascin-X suggests a role in limb, muscle, and heart development. Dev Dyn 1995, 203, (4), 491-504. 12. Egging, D. F.; van Vlijmen, I.; Starcher, B.; Gijsen, Y.; Zweers, M. C.; Blankevoort, L.; Bristow, J.; Schalkwijk, J., Dermal connective tissue development in mice: an essential role for tenascin-X. Cell Tissue Res 2006, 323, (3), 465-74. 13. Chiquet-Ehrismann, R.; Hagios, C.; Schenk, S., The complexity in regulating the expression of tenascins. Bioessays 1995, 17, (10), 873-8. 14. Erickson, H. P., Tenascin-C, tenascin-R and tenascin-X: a family of talented proteins in search of functions. Curr Opin Cell Biol 1993, 5, (5), 869-76. 15. Elefteriou, F.; Exposito, J. Y.; Garrone, R.; Lethias, C., Binding of tenascin-X to decorin. FEBS Lett 2001, 495, (1-2), 44-7. 16. Elefteriou, F.; Exposito, J. Y.; Garrone, R.; Lethias, C., Cell adhesion to tenascin-X mapping of cell adhesion sites and identification of integrin receptors. Eur J Biochem 1999, 263, (3), 840-8.  82  17. Zweers, M. C.; van Vlijmen-Willems, I. M.; van Kuppevelt, T. H.; Mecham, R. P.; Steijlen, P. M.; Bristow, J.; Schalkwijk, J., Deficiency of tenascin-X causes abnormalities in dermal elastic fiber morphology. J Invest Dermatol 2004, 122, (4), 885-91. 18. Burch, G. H.; Gong, Y.; Liu, W.; Dettman, R. W.; Curry, C. J.; Smith, L.; Miller, W. L.; Bristow, J., Tenascin-X deficiency is associated with Ehlers-Danlos syndrome. at Genet 1997, 17, (1), 104-8. 19. Schalkwijk, J.; Zweers, M. C.; Steijlen, P. M.; Dean, W. B.; Taylor, G.; van Vlijmen, I. M.; van Haren, B.; Miller, W. L.; Bristow, J., A recessive form of the Ehlers-Danlos syndrome caused by tenascin-X deficiency. ew England Journal of Medicine 2001, 345, (16), 1167-1175. 20. Elefteriou, F.; Exposito, J. Y.; Garrone, R.; Lethias, C., Characterization of the bovine tenascin-X. Journal of Biological Chemistry 1997, 272, (36), 22866-22874. 21. Jones, F. S.; Hoffman, S.; Cunningham, B. A.; Edelman, G. M., A detailed structural model of cytotactin: protein homologies, alternative RNA splicing, and binding regions. Proc atl Acad Sci U S A 1989, 86, (6), 1905-9. 22. Erickson, H. P.; Inglesias, J. L., A six-armed oligomer isolated from cell surface fibronectin preparations. ature 1984, 311, (5983), 267-9. 23. Spring, J.; Beck, K.; Chiquet-Ehrismann, R., Two contrary functions of tenascin: dissection of the active sites by recombinant tenascin fragments. Cell 1989, 59, (2), 325-34. 24. Oberhauser, A. F.; Marszalek, P. E.; Erickson, H. P.; Fernandez, J. M., The molecular elasticity of the extracellular matrix protein tenascin. ature 1998, 393, (6681), 181-185. 25. Bork, P.; Doolittle, R. F., Proposed acquisition of an animal protein domain by bacteria. Proc atl Acad Sci U S A 1992, 89, (19), 8990-4. 26. Rief, M.; Gautel, M.; Schemmel, A.; Gaub, H. E., The mechanical stability of immunoglobulin and fibronectin III domains in the muscle protein titin measured by atomic force microscopy. Biophysical Journal 1998, 75, (6), 3008-3014. 27. Oberhauser, A. F.; Badilla-Fernandez, C.; Carrion-Vazquez, M.; Fernandez, J. M., The mechanical hierarchies of fibronectin observed with single-molecule AFM. Journal of Molecular Biology 2002, 319, (2), 433-447. 28. Rief, M.; Gautel, M.; Oesterhelt, F.; Fernandez, J. M.; Gaub, H. E., Reversible unfolding of individual titin immunoglobulin domains by AFM. Science 1997, 276, (5315), 1109-1112. 29. Carrion-Vazquez, M.; Oberhauser, A. F.; Fowler, S. B.; Marszalek, P. E.; Broedel, S. E.; Clarke, J.; Fernandez, J. M., Mechanical and chemical unfolding of a single protein: A comparison. Proceedings of the ational Academy of Sciences of the United States of America 1999, 96, (7), 3694-3699. 30. Zweers, M. C.; van Vlijmen-Willems, I. M.; van Kuppevelt, T. H.; Mecham, R. P.; Steijlen, P. M.; Bristow, J.; Schalkwijk, J., Deficiency of tenascin-X causes abnormalities in dermal elastic fiber morphology. Journal of Investigative Dermatology 2004, 122, (4), 885-891. 31. Batey, S.; Clarke, J., Apparent cooperativity in the folding of multidomain proteins depends on the relative rates of folding of the constituent domains. Proceedings of the ational Academy of Sciences of the United States of America 2006, 103, (48), 18113-18118. 32. Krammer, A.; Lu, H.; Isralewitz, B.; Schulten, K.; Vogel, V., Forced unfolding of the fibronectin type III module reveals a tensile molecular recognition switch. Proceedings of the ational Academy of Sciences of the United States of America 1999, 96, (4), 1351-1356. 33. Gao, M.; Craig, D.; Vogel, V.; Schulten, K., Identifying unfolding intermediates of FNIII10 by steered molecular dynamics. Journal of Molecular Biology 2002, 323, (5), 939-950.  83  34. Gao, M.; Craig, D.; Lequin, O.; Campbell, I. D.; Vogel, V.; Schulten, K., Structure and functional significance of mechanically unfolded fibronectin type III1 intermediates. Proceedings of the ational Academy of Sciences of the United States of America 2003, 100, (25), 14784-14789. 35. Oberhauser, A. F.; Marszalek, P. E.; Carrion-Vazquez, M.; Fernandez, J. M., Single protein misfolding events captured by atomic force microscopy. ature Structural Biology 1999, 6, (11), 1025-1028. 36. Wright, C. F.; Teichmann, S. A.; Clarke, J.; Dobson, C. M., The importance of sequence diversity in the aggregation and evolution of proteins. ature 2005, 438, (7069), 878-881. 37. Lethias, C.; Carisey, A.; Comte, J.; Cluzel, C.; Exposito, J. Y., A model of tenascin-X integration within the collagenous network. FEBS Lett 2006, 580, (26), 6281-5. 38. Cao, Y.; Li, H. B., Polyprotein of GB1 is an ideal artificial elastomeric protein. ature Materials 2007, 6, (2), 109-114.  84  Chapter 3 Investigation of the anomechanical Properties of a on-Mechanical Protein: Bacillus Circulans Xylanase 3.1 Introduction to Bacillus Circulans Xylanase 3.1.1 BCX is a Bacterial Enzyme Responsible for the Breakdown of Cell Walls Cellular walls within plants provide a tough yet flexible layer that gives the surrounding tissue rigidity and offers protection against mechanical stress. Plant cell walls may be broken down into two components: the primary cell wall, a thin and flexible layer next to the cellular membrane and the lignin-rich secondary cell wall that forms a more rigid, waterproof layer. The predominant component within the primary cell wall is cellulose, hemicellulose and pectin. The cell wall matrix is assembled out of linked networks of these polysaccharides that imparts high tensile strength and elasticity to tissues, but is also difficult to hydrolyze. This makes it challenging to break down and digest many forms of plant tissues, a process that is unique to organisms that produce specific enzymes designed for the hydrolysis of these durable assemblies. These organisms are generally communities of anaerobic micro organisms that exist in a symbiotic relationship with a host who benefits from the breakdown of otherwise indigestible plant material. That mammals are able to digest plant material at all hinges on the action of the anaerobic bacteria, fungi and protozoa lining the gut which are responsible for the degradation of polysaccharides plant material ingested. Other organisms, such as arthropods and gastropods, also produce glycosidase hydrolase enzymes in the production of monomeric xylan critical to cell metabolism.1 These organisms typically produce cellulase and xylanase enzymes in tandem, secreting them into the ECM to hydrolyze long chain cellulose and hemicellulose too large to penetrate into the cell. Glycoside hydrolases usually exhibit a modular construction made of conserved catalytic domains and carbohydrate binding modules (CBM) that are joined by flexible linker sequences.2-4 Domains within these proteins are typically structurally independent and exhibit some degree of functional autonomy.5-7 Cleavage of polysaccharide substrates is accomplished via substrate binding to the CMB regions, which allows for subsequent association of catalytic domains as well as interference with the substrate structure.8, 9  85 A version of this chapter will be submitted for publication Jollymore, A., Withers, S.W., Lawrence, L.P., Li, H. (2009) The Folding Behavior of Bacillus Circulans Xylanase Revealed Using Single Molecule AFM  3.1.2 Variability within the Structure of Xylan Has Resulted in a Multiplicity of Xylanase Forms Bacillus circulans xylanase (BCX) is a retaining endo-1,4-xylanase from the glycoside hydrolase family 11 clan GH-C responsible for the hydrolysis of xylans.10, 11 Xylans are the second most common polysaccharide found within plant biomass, and the most common hemicellulose polysaccharide located primarily within the secondary cellular walls of land plants; it is found to a lesser degree within the primary cell wall.12-14 Xylans in plants are interspersed with and covalently bonded to an overlying sheath of lignin, coating underlying strands of cellulose via the formation of hydrogen bonds.13-16 This arrangement is thought to be important for fibre cohesion and the integrity of the plant cell wall.17 The backbone of xylan is formed by β-(1→4) linked β-xylopyranose residues. Xylan is a complex polysaccharide, usually exhibiting some degree of backbone substitution and a highly branched structure whose composition varies dependent on the plant species. Linear unsubstituted forms of xylan are actually relatively rare in comparison to the plethora of heteropolysaccharide xylan forms, and are encountered only in certain species such as esparto grass,18 tobacco plants,19 and some species of algae.20, 21 The degree of backbone substitution varies and may include the substitution of glucuronopyranosyl, 4-O-methyl-D-glucuronopyranosyl, α-L-arabinofuranosyl, acetyl, feruloyl and p-coumaroyl side chain groups.12, 22, 23 The size of the polymerized xylan chain is also variable depending on the xylan type, ensuring huge variation in naturally occurring xylans. For example, xylans from hardwood species exist as O-acetyl-4-O-methylglucuronoxylan polymerized into 150-200 β-xylopyranose residues, where softwood xylans subsist as arabino-4O-methylglucuronoxylan polymerized into shorter chains of 70-130 residues.12 This multiplicity of xylans has resulted in the evolution of a large variety of xylanases that exhibit a wide range of activity and binding specificity. Xylanases are typically classified according to the primary sequence of their catalytic domain, with family members exhibiting analogous structural features and catalytic activity.24 Family 11 xylanases such as BCX are characterized by their low molecular weight, basic pI, as well as comparable quaternary structure and molecular functionality.  86  Figure 3.1 The Xylan Backbone: The Substrate for BCX  α-4-O-methylglucuronic acid  endo-1,4-β-xylanase  acetylxylan esterases  α-L-arabinofuranosidase  α-D-glucuronidase  β-xylose chain  Β-D-xylosidase  feruloyl and p-coumaroyl esterases  α-L-arabinofuranosyl feruloyl and p-coumaroyl side chains Figure 3.1 The structure of xylans and specific xylanase cleavage sites. The xylan backbone is composed of linked 1,4-β-xylose residues. Several typical branching sidechains are shown above (bold italic type; Ac=acetyl groups). The variety of xylans encountered in nature has resulted in a plethora of xylanases capable of hydrolyzing xylans at a multiplicity of sites and branching sidechains. Figure adapted from [23]  3.1.3 Investigating Xylanases as a ‘Greener’ Bleaching Agent within Paper Industries Xylanases have been the subject of much interest and subsequent research due to their potential application to industrial processes that involve the decomposition of plant materials. The pulp and paper industry in particular has devoted a large amount of time attempting to integrate xylanases into the processing of wood pulp into paper. Highly alkali conditions used to extract brown-coloured lignin during the pulping process cause the precipitation of short-chain xylans into a crystalline layer onto cellulose microfibrils. This layer prevents the full extraction of lignin, resulting in a decrease in pulp brightness.25 Improving the colour of the pulp typically requires harsh bleaching steps that use environmentally destructive chlorine-containing compounds that are difficult to dispose of. The interest in xylanases stems from the desire to implement an enzymatic stage that would break down the crystalline xylan barrier, allowing for the use of less bleaching agent in a more environmentally friendly process. To this end, xylanases have been demonstrated on a semi-industrial scale in the dissolution of pulp, yielding 87  cellulose for the production of rayon, and bio-bleaching of wood pulps.26 Xylanase use is limited by a number of factors. The harsh conditions under which pulping is accomplished means that enzymes must be optimized for maximal efficiency within a highly alkaline, high temperature environment, making necessary the use of extremophilic xylanases. In addition, the large scale use of enzymes is a more cost intensive process than traditional methods, limiting the industrial use of xylanases despite its potential as a ‘greener’ bleaching agent. 3.1.4 BCX Structure Like other Family 11 xylanases, BCX exhibits a β-sandwich structure that has been extensively characterized using X-Ray crystallography and NMR spectroscopy.27-29 The structure forms a 20 kDa β-sandwich that curls into a shape reminiscent of a partially closed right hand.30 Within this, β-pleated sheets curve into a hydrophobic pocket surrounding the active site cleft which is bordered on one side by an extended flexible sequence (the ‘thumb’ of the hand).29, 31 BCX is predominantly composed of β-sheets, with a single α-helix packed against the underside of one β-sheet.  88  Figure 3.2 The Structure of Bacillus Circulans Xylanase (BCX)  Figure 3.2 The structure of BCX as solved using X-Ray Crystallography. BCX is a single domain family 11 xylanase protein approximately 20 kDa in mass. BCX structure forms a β-sandwich shape that curls in upon itself in a shape reminiscent of a partially closed right fist, where the active site is located in the palm. The N and C terminus are located within close proximity of each other on the top of the molecule as shown.  3.1.5 BCX Mechanism: How Xylan Cleavage is Accomplished The structure exhibited by BCX is finely tuned in order to hydrolyse long chains of xylans. BCX has been extensively investigated in terms of this mechanism, where the specific structural adaptations used to bind and cleave xylans, the specific kinetics of xylan binding, as well as the functional effects of pH have been evaluated.32-36 BCX binds xylan chains nonspecifically anywhere along the length of the chain, preferentially binding chains composed of at least three xylan monomers.30 Thus, xylan cleavage by BCX results in the production of varying sizes of cleaved xylans, including monomeric or dimeric xylan. BCX, like most family 11 xylanases, does not contain a carbohydrate binding module (CBM), but instead binds xylan within the active site cleft as well as at a non-catalytic site located at the surface of the BCX 89  structure.30 This secondary binding site is a shallow groove that is lined with exposed polar sidechains, functioning cooperatively with the active site to increase xylan binding affinity and catalysis, acting as a pseudo CBM.30 BCX, as well as other members of family 11 glycosidases, cleaves xylans with a double displacement mechanism that results in an overall retention of anomeric conformation. 10 The first stage of this mechanism is a glycosylation step that forms a covalent intermediate; this species is subsequently hydrolysed during the deglycosylation stage via oxocarbenium-ion-like transition states.37-39 The initial glycosylation step is initiated by the nucleophilic attack of the residue Glu78 upon the glycosidic bond of the xylose polymer bound within the active site of the protein. The formation of a covalent glycosyl-enzyme intermediate, where the proximal saccharide is distorted into the 2,5B conformation,28 is completed by the protonated active site residue Glu172, which acts as a general acid by donating a proton to the departing aglycon group.28, 40, 41 The pKa of these two critical active site residues reflects this mechanism. The pKa of the nucleophilic glutamate is much lower than that of the acidic Glu172 (pKa=4.6 and 6.7 respectively).42 BCX functions optimally at a pH of 5.7, ensuring the correct initial protonation states for both of these catalytic residues.40 In the second stage of xylan hydrolysis, the deprotonated residue Glu172 (pKa 4.2) acts as a general base, hydrolyzing the glycosyl intermediate to its final cleaved state.  90  Figure 3.3 BCX Cleaves Xylans Through A Double Displacement Mechanism  A  C  pKa 6.7 Glu172  Glu172  δ HO  O  O  O H  O RO  RO  GLYCOSYLATIO OR'  HO  O OR'  δ  HO  OH  OH Oδ  O  O  O  Glu78 Glu78 pKa 4.2  pKa 4.6  Glu172 -R'OH O  Glycosyl-Enzyme Intermediate  RO  O  H  O  HO  O OH O  H O  pKa 6.7 Glu172 Glu 78  B  Glu 172  HO  O  δO O RO  RO OH  HO  O  O  δ  HO  OH  DEGLYCOSYLATIO  OH  OH Oδ  O  Glu78 pKa 4.6  O  H  O  Glu 78  Figure 3.3 Xylan cleavage by BCX. Xylan binds within the active site cleft of BCX, where two residues located within the active site are critical in the cleavage of xylan substrates (Figure 3.3 A; Glu78 in red, Glu172 in blue). A secondary xylan binding site located outside of the active site has recently been demonstrated to increase the binding affinity of BCX to long chain xylans (Figure 3.3 B, residues participating in xylan binding in red; adapted from Ref [30]). Xylans are cleaved by BCX via a double displacement mechanism (Figure 3.3 C). In the initial glycosylation step, the low pKa of Glu78 allows it to act as a nucleophile as it attacks the β-(1-4)-linked xylose chain. Glu172 acts as a general acid catalysis to form the glycosyl-enzyme intermediate. Deglycosylation of this intermediate occurs when Glu172 acts as a general base (pka=4.2) to hydrolyse the glycosyl-enzyme intermediate. Figure 3.3 C adapted from Ref [33].  3.2 BCX as a Force Spectroscopy Model System Bacillus circulans xylanase clearly demonstrates non-mechanical functionality. Nothing about the catalytic functionality, structure, or physiological environment suggests that force plays anything more than an incidental role in the operation of BCX. The mechanical properties exhibited by BCX remain unknown despite the large body of information already elucidated. To 91  this end, BCX has been extensively studied in terms of its structure, mechanism, binding kinetics and binding partners by a variety of non-force techniques providing for the opportunity to add and compare to this details about the mechanical properties of BCX. Investigating these features also allows for the contrast to be made between BCX as a well-defined non-mechanical protein and other glycosidases such as T4 lysozyme as well as mechanical proteins such as tenascin-X. These comparisons will aid in the determination of molecular characteristics involved in mechanical stability. 3.2.1 Previous Studies on the Thermal Stability of BCX Previous studies on BCX have elucidated the thermal stability of the wild type protein as well as a number of active site and disulfide bonded mutants. Investigating the thermal stability of BCX using differential scanning calorimetry revealed the high pH dependence of wild type stability, with a ~3°C difference in Tm for wild type BCX between a pH of 5 and a pH of 8.42 Certain point mutations within the structure of BCX were also found to increase the thermal stability of BCX; replacing Glu172 with an uncharged cysteine was found to increase the tm by 7.8°C at pH 8, an effect that was ascribed to the diminished electrostatic repulsion between nearby carboxylate groups upon the insertion of an uncharged residue into the active site.42 The insertion of disulfide bonds into the structure of BCX has previously revealed several BCX mutants who show greater thermal stability than that of the wild-type protein.43 Thermal denaturation of BCX and its mutants revealed the apparent irreversibility of this transition. BCX proteins that have been thermally unfolded do not exhibit any evidence that refolding occurs upon cooling and subsequent reheating, making exposition of folding kinetics impossible by these methods. Using the AFM to probe BCX unfolding represents a method of determining whether irreversible unfolding is a property of the protein or of the method undertaken to denature the protein, as well as determining whether mutations that result in an increase in thermal stability also increase mechanical stability. Previous study of BCX has also resulted in the discovery of a number of inhibitor compounds that mimic the glycosyl-enzyme intermediate bound in the active site of BCX in the hydrolysis of xylan. These compounds include the covalent inhibitor 2’,4’-dinitrophenyl 2deoxy-2-fluoro-β-xylobioside, which acts to block the active site of BCX by forming a long 92  lived glyosyl-enzyme intermediate.41 30 This effectively freezes the protein into a bound conformation, allowing for the elucidation of the dynamics and conformational changes that occur as the enzymatic mechanism proceeds using NMR spectroscopy and X-ray crystallography.28, 44 How the binding of substrates affects the mechanical stability of BCX has not been previously investigated. However, the effect that substrate binding has on the mechanical stability of proteins has been previously investigated for other enzymatic systems. These studies seemingly indicate that binding has negligible effects on apparent mechanical stability.45 The effect substrate binding has upon the mechanical properties of BCX has not been previously investigated, but is possible to evaluate due to the existence of long-lived inhibitors that mimic the transition state of BCX.  93  Figure 3.4 Inhibitor Binding Forms a Long-Lived Covalent Species pKa6.7 Glu 172  HO  O  OH O HO HO  NO2  O O  HO  O  F  O  O NO2  Glu 78  pKa 4.6  pKa 4.2 Glu 172  O  NO2  O  OH O HO HO  O  O O  HO F O  NO2  O Glu 78  Figure 3.4 Inhibitor binding to BCX and the formation of a long lived covalently bound species that mimics the transition state. Inhibitor binding to BCX halts hydrolysis after the formation of the glycosyl-enzyme intermediate. Nucleophilic attack of Glu78 occurs as in the case for xylan substrates, and is facilitated by the formation of the 2,4-dinitrophenyl leaving group. Further hydrolysis of the intermediate is slowed by the fluorine present at position 2 on the proximal saccharide. Figure adapted from Ref[28].  The thorough structural investigation of BCX by both NMR and X-Ray crystallography makes it an attractive force spectroscopy model protein. This technique allows for the elucidation of specific structural elements that contribute to mechanical stability through the eventual use of steered molecular dynamics (SMD), as well as allowing for the structural comparison between the mechanical properties of proteins previously determined using single molecule AFM. SMD, coupled with unfolding information gleaned from single molecule experiments, can provide detailed insight into how unfolding and folding of a protein occurs. These techniques have been extensively used in tandem with AFM to provide information about the molecular mechanisms behind folding and unfolding. Furthermore, uncovering residues that 94  form bonds key to mechanical stability helps illustrate potential intermediates formed during the folding or unfolding process, and allows for determination of possible effects as a result of point mutations. Prior knowledge of the structure of a protein is critical in order for a high resolution simulation; this presents a bottleneck in the case of bovine tenascin-X due to a lack of structural information about this form of tenascin. Knowing the structure of BCX not only allows for the simultaneous use of SMD in order to elucidate the experimental mechanical information gained from AFM, it also allows for the mechanical comparison to be made on the basis of protein structure. Certain trends in mechanical characteristics have begun to emerge for specific structural elements contained within proteins. As BCX is a non-mechanical protein with well defined structure, it makes an appealing comparative model system in the interests of expanding how molecular characteristics determine observed mechanical properties. 3.3 Investigating the Mechanical Properties of BCX 3.3.1 Research Overview The following single molecule AFM analysis of BCX focused first on the mechanical unfolding and folding of a protein construct containing wild type BCX. The effect of binding this BCX-containing construct to the covalent inhibitor 2’,4’-dinitrophenyl 2-deoxy-2-fluoro-βxylobioside was also investigated using single pull experiments. Finally, three disulfide bond mutants of BCX were investigated to determine the effects such bonding would have on the mechanical properties in comparison to wild type BCX. One of these mutants, BCX100x148, was singled out in particular due to its proven increase in thermal stability in comparison to the wild type protein. 3.3.2 Investigating the Mechanical Properties of BCX using Single Molecule AFM In order to determine the mechanical properties of BCX, a polypeptide construct was made with one BCX bracketed from both ends by four repeats of the molecular fingerprint protein GB1. This was done in a similar manner as accomplished in the previous AFM investigation of T4 lysozyme.46 Having eight repeated monomers of GB1 within the protein construct does diminish the number of BCX unfolding events observable within a single force extension curve, yet was done to take advantage of several of GB1’s characteristics. GB1 is 95  easily expressed within the E.coli cell lines utilized within this work, and demonstrates a high degree of AFM ‘friendliness’, being easily adsorbed onto glass and subsequently picked up with a silicon nitride tip. Its presence also allows for the unambiguous determination of single molecule unfolding events by examination of the resultant force extension curve.  96  Figure 3.5 The BCX Polyprotein and its Resultant Force Extension Fingerprint  A  B  64 nm  64 nm  18 nm  200 pN 50 nm  Figure 3.5 The force extension traces of a polyprotein containing BCX. The polyprotein construct investigated using single molecule AFM contained one BCX domain bracketed by 4 respective repeats of GB1 (Figure 3.5 A). GB1 served as a nanomechanical fingerprint in the identification of single molecule unfolding events observed with the force extension curves of the [GB1]4BCX[GB1]4 polyprotein (Figure 3.5 B, blue fits). Unfolding events corresponding to the unfolding of BCX occurred at low extension, and were fit using a Worm-Like Chain model of polymer elasticity (red lines). Most events occurred approximately 60 nm from the first GB1 unfolding event, yet some variation was exhibited during the fitting of BCX unfolding using the WLC model (lower two traces). As well, the formation of dimers was noted in curves that exhibited more than one BCX unfolding event (upper two traces).  The mechanical unfolding of a (GB1)4BCX(GB1)4 resulted in a force extension curve that demonstrated a typical sawtooth pattern of unfolding. The presence of both GB1 and BCX domains within the unfolding of the (GB1)4BCX(GB1)4 polyprotein was verified by the presence of unfolding events that could be fit using two separate values of ∆Lc. Unfolding events were 97  observed that could be fit to a ∆Lc of ~18 nm, corresponding to the unfolding of GB1 domains within the stretched polyprotein. These events tended to occur at high extension towards the end of the force extension curve (Figure 3.5, blue Worm-like chain fits). Based on the number of GB1 repeats within the polyprotein construct, the maximum number of GB1 unfolding events within the force extension of (GB1)4BCX(GB1)4 should be eight, knowing that the polyprotein is picked up randomly along its contour during AFM force extension experiments. This number was regularly exceeded, corresponding to the polymerization of (GB1)4BCX(GB1)4 into dimers through the cysteine residues engineered at the C terminus (Figure 3.5 B, upper two force extension traces). The number of GB1 unfolding events observed within a single force extension trace was as low as five and as high as 15. A second distribution of unfolding events could be fit with values of ∆Lc approximately equal to 64 nm. These events tended to occur at lower extension than that of GB1 and were ascribed to the unfolding of BCX within the (GB1)4BCX(GB1)4 structure (Figure 3.5, red Worm-like chain fits). The formation of dimers assembled out of two (GB1)4BCX(GB1)4 was also made apparent by counting the number of BCX unfolding events within the (GB1)4BCX(GB1)4 force extension curve: as there is only one BCX domain present within the engineered protein examined, the presence of two unfolding events occurring with a ∆Lc of ~64 nm signified the formation of (GB1)4BCX(GB1)4 dimers. Initially, the unfolding of the BCX-containing polyprotein was attempted using a single pull protocol, where a single molecule of protein is stretched between the tip and surface until it detaches from either of these surfaces. Subsequent single molecule measurements are then made by picking up a second molecule and repeating the procedure. Because of the low extension at which BCX unfolds as well as the apparently low unfolding force of BCX, such a protocol made it difficult to differentiate unfolding peaks of BCX from non-specific unfolding events that occur at low extension. Thus, subsequent experiments were done in a refolding fashion, where a single molecule of protein was extended, allowed to refold upon relaxation, and re-extended. All refolding experiments were done at a constant pull rate of 400 nm/s. Effects stemming from heterogeneity between BCX molecules were circumvented by examining a large number of refolding experiments on different molecules of protein. The elastic nature of BCX is revealed in these experiments, in that a molecule of protein containing BCX may be unfolded and refolded a number of times without apparent change in its mechanical properties.  98  Figure 3.6 Force Histogram and Contour Length Increment of BCX  A  No. of Events  30 25 20 15 10 5 0 0  50  100  150  200  Force (pN)  B  C 1600  80  1400  No. of Events  No. of Events  1200 1000  60  40  800 600 400 200 0 16  20  18  20  22  24  ∆Lc (nm) 0 40  50  60  70  80  90  ∆Lc (nm)  Figure 3.6 The average unfolding force and ∆Lc of BCX. The unfolding force exhibited by BCX demonstrates a large range with an average of 55 ± 30 pN (Figure 3.6 A). The force histogram of BCX unfolding events indicates that the mechanical stability of BCX is relatively low. The contour length increment of BCX is broad, reflecting variation observed in force extension curves (Figure 3.6 B). The observed ∆Lc of 64 ± 8 nm (n=260) reflects the number of amino acids participating in the fold of BCX. The broadness of this distribution is in direct contrast with the distribution of ∆Lc determined from GB1 unfolding events within the same force extension events (Figure 3.6 C). This distribution is narrow, centered about 17.5 ± 1.0 nm, and corresponds well with the known ∆Lc of GB1.  Compiling the unfolding events caused by the refolding of (GB1)4BCX(GB1)4 on the basis of their unfolding force resulted in the creation of an unfolding force histogram specifying the average unfolding force of BCX. This was done by examining the force extension curves of a large number of unfolding events from a large number of single molecules unfolded using a refolding protocol. Because of the low-extension unfolding of BCX, care was taken to ensure that unfolding events were taken only from experiments whose cleanliness was verified by the  99  absence of such non-specific unfolding events. Pooling these stringently discriminated BCX unfolding events resulted in an average unfolding force for BCX of 55 ± 30 pN (n=250). Low extension unfolding events corresponding to the unfolding of BCX was fitted using the Worm-Like Chain model of polymer extensibility. This allowed for the measurement of the contour length increment (∆Lc) of BCX. A value of ~0.5 nm for the persistence length was used in the fitting the unfolding events of BCX. The contour length increment of BCX was found to be 64 ± 8 nm (n=260). This corresponds fairly well to the determined domain size of 185 amino acids and an N-C terminal distance of ~0.5 nm ( Lc= (185 a.a × 0.36 nm/a.a)-0.5 nm). However, the contour length increment measured for BCX broadens about this central value of ∆Lc. Unlike proteins such as T4 lysozyme, where the spread of ∆Lc was noted to be the result of well-defined unfolding intermediate states,46 the variation present within BCX is less quantifiable, occurring with less predictability than that of T4 lysozyme.  100  3.3.2.1 The Pull Rate Dependence of BCX Determines its Unfolding Rate Parameters Figure 3.7 Unfolding Force Dependence on Pulling Speed: Unfolding Kinetics  120  Unfolding Force (pN)  100 80 60 40 20 0 5 6 7 8 9  100  2  3  4  5  6 7 8 9  2  3  4  5  1000  Pull Speed (nm/s) Figure 3.7 The dependence of unfolding force on pulling speed allows for the elucidation of unfolding kinetics. BCX exhibits a relatively weak relationship between the unfolding force observed and the rate at which the protein is pulled. Fitting this dependence using a Monte Carlo simulation (solid line Figure 3.7) -2 -1 allowed for the determination of the unfolding rate constant (αo=3×10 s ) and unfolding distance (∆xu=0.63 nm).  The dependence of observed unfolding force is a property intrinsic to the manner in which AFM unfolding of a single molecule is done; thus, the unfolding kinetic parameters of BCX may be determined by this dependence as previously discussed. The construct (GB1)4BCX(GB1)4 was pulled at speed varying from 50 nm/s to 5000 nm/s. Based on the contour length increment exhibited by BCX within refolding experiments done at 400 nm/s, BCX peaks were those that exhibited a contour length increment of ~64 nm. The average unfolding force of these peaks were then compared on the basis of the speed at which unfolding was observed (Figure 3.7). The dependence of force on pulling speed was small, and was fit using a Monte Carlo simulation in order to determine ∆xu and αO as previously described. The pull rate dependence of BCX was fit using a unfolding distance ∆xu of 0.63 nm and a unfolding rate constant αO of 3×10-2 s-1.  101  3.3.2.2 Refolding of BCX Reveals Folding Kinetics In order to determine the kinetic parameters that described the folding behavior of BCX, a double pulse protocol was utilized as previously described. Briefly, this involved the complete unfolding of a single molecule of protein by extending it to the point just before detachment occurs and determining the number of BCX molecules present within the polyprotein,  total.  The  protein was then relaxed back to zero extension and allowing it to refold in a varying amount of time ∆t. After this time passes, the molecule was re-extended to determine whether BCX had refolded during this time, where the number of unfolding events equals refold/ total  refold.  The ratio  at a number of refolding times ∆t may be fit to the single exponential distribution  described by:  refold  (  = A 1 − e − βt  )  (3.1)  total  where  refold/ total  is the folding probability of BCX domains at a relaxation time t, and β refers  to the folding rate constant. As this was done using a construct containing one BCX repeat, the probability of observing refolding after unfolding is either 0% or 100%, where a minority of curves from dimerized polyproteins exhibited two BCX unfolding events. Compiling a large number of refolding curve pairs resolves the probability of observing refolding after ∆t, resulting in the distribution observed in Figure 3.8. The refolding kinetics of BCX was fit using a single exponential distribution with a folding rate constant β of 0.56 s-1. This study represents the first successful elucidation of BCX folding kinetics despite previous attempts using ensemble denaturation methods, illuminating the utility of single molecule AFM techniques in conducting experiments impossible to conduct under ensemble conditions.  102  Figure 3.8 The Folding Kinetics of BCX  1.0  Folding Probability  0.8 0.6 0.4 0.2 0.0 0  1  2  3  4  ∆t(s) Figure 3.8 The folding kinetics of BCX. The folding behavior of BCX was determined using a refolding double pulse method, in which a single molecule of BCX containing polyprotein was unfolded, relaxed and allowed to refold during the time ∆t, and re-extended to determine whether BCX could refold during ∆t. As the [GB1]4BCX[GB1]4 polyprotein contained only one BCX domain, statistics were generated by comparing refolding traces at a variety of ∆t values. The folding probability of BCX was fit using a single -1 exponential distribution with folding rate constant β of 0.56 s .  3.3.2.3 The Unfolding of Wild Type BCX Reveals its anomechanical Behavior Forcibly unfolding BCX using single molecule AFM reveals a number of features not previously elucidated. Unfolding the polyprotein construct (GB1)4BCX(GB1)4 reveals typical sawtooth unfolding behaviour, in which peaks of GB1 are easily identified from those stemming from the unfolding of the single BCX module within the structure of the protein. The unfolding of BCX occurs at low extension, approximately 60 nm from the first GB1 unfolding event. Mechanical unfolding of BCX using a refolding protocol, as was done to aid the identification of BCX unfolding events, reveals the propensity of BCX to refold after unfolding without apparent change in its mechanical properties. That BCX undergoes a reversible force unfolding transition is in direct contrast to the observed irreversible nature of thermal unfolding, in which ensembles of BCX molecules denatured using high temperatures remain unfolded after removing from heat. This apparent differentiation in unfolding behaviour may be a result of the low concentration in which AFM experiments are done, preventing the aggregation of protein molecules previously 103  noted within ensemble experiments. This aggregation was postulated to be a cause for the irreversibility of thermal unfolding, and would be unlikely to occur within the low protein concentrations used within AFM experiments. The reversibility of mechanical unfolding allows for the elucidation of folding properties using single molecules, properties that have previously been impossible to examine using techniques that result in irreversible unfolding of BCX. Examining the magnitude of the low extension BCX unfolding event reveals its average unfolding force. This protein unfolds at low applied force, signifying a lower degree of mechanical stability than either GB1 or tenascin-X as examined beforehand within this document. This reduction is significant, with BCX unfolding occurring at ~100 pN lower than that of the molecular fingerprint GB1. This low apparent mechanical stability is not surprising considering the intrinsic functionality that BCX demonstrates: unlike mechanical proteins such as tenascin, mechanical stability is apparently unnecessary for BCX efficacy and does not play a large role within its physiological environment. The average unfolding force is similar to that exhibited by T4 lysozyme (~50 pN),46 another protein whose intrinsic functionality is divorced from apparent force. BCX exhibits a drastically different structure than T4 lysozyme, which exists as a 164 residue protein composed of 10 α-helices and 4 short β-strands.47 Proteins with a large amount of α-helical content have previously demonstrated a tendency towards lowered mechanical stability; thus, it is not surprising that the predominantly α-helical T4 lysozyme unfolds at low applied force. This is the same case for the unfolding of the globular RNase protein barnase, a small non-mechanical protein composed of 110 amino acids. Its structure is dominated by a hydrophobic core formed by the packing of an N-terminal α-helix against a five stranded β-sheet, along with the formation of two smaller hydrophobic pockets formed by the packing of the second and third α-helix against the same β-sheet.48 Thus, barnase is composed equally of 20% β-strands and 20% α-helices, with the remainder of the protein existing as unstructured sequences. The average unfolding force of barnase was noted to be ~70 pN, reflecting the decreased mechanical stability of this protein in comparison to Ig domains or any of the tenascins examined to date.48 Barnase exhibits an increased average unfolding force when compared to that of T4 lysozyme, consistent with trends that ascribe higher mechanical stability to proteins with increased β-strand content.49 This trend is discontinuous for BCX, which is constituted predominantly of β-sheets folded against one another, displaying the presence of only  104  one α-helix and one unstructured loop region. Proteins with increased β- strand content tend to reveal a higher degree of mechanical stability;49 therefore, it is surprising that the mechanical stability of BCX is similar to that of T4 lysozyme and smaller than that of barnase. Simply ascribing mechanical stability to global trends in the type of structure exhibited cannot be broadly applied across a range of protein structures in order to predict mechanical stability. In the case of BCX, low mechanical stability may be a result of the manner in which force is applied to the structure of the protein. Applied mechanical force that causes ‘zippering’ rather than ‘shearing’ unfolding behaviour has previously been ascribed to lowered mechanical stability.50, 51 Modeling the behaviour of BCX as it is forcibly unfolded using steered molecular dynamics or other such method would be an excellent approach towards determining whether this is the case. The low extension peak within the force extension curve of (GB1)4BCX(GB1)4 corresponding to the unfolding of BCX could be fit to the higher extension unfolding of GB1 using the Worm Like Chain. Typically, the distribution of ∆Lc values used to fit protein unfolding events is relatively narrow and represents the number of amino acids participating in folding. BCX did not exhibit the same fixed increment within its exhibited contour length as is typical for protein unfolding studies, exhibiting a larger distribution about an average ∆Lc that corresponds to the theoretical contour length. This was the same case in the unfolding of barnase, which exhibited a similar variation in values of ∆Lc.48 Such deviation was also present within the observed ∆Lc during the unfolding of T4 lysozyme; unlike barnase or BCX, ∆Lc deviations occurred in a quantifiable pattern. These arrangements signified unfolding intermediate states formed and provided a description of partitioned kinetic pathways ascribed as the protein unfolds.46 Though the ∆Lc observed within the unfolding of BCX were less quantifiable within specific blueprints as for T4 lysozyme, it is possible that contour length increment variability is due to a similar mechanism. If this is so, BCX details a number of pathways as it unfolds, resulting in the formation of a number of unfolding intermediate structures in a more complex and less definable manner than that of T4 lysozyme. The apparent variation present within ∆Lc may be due to the unfolding of BCX domains that are either partially unfolded prior to the application of force using single molecule AFM or effects that occur due to possible associations occurring between BCX and closely coupled domains of GB1.  105  Previously utilized thermal denaturation results in the irreversible unfolding of BCX;34, 42, 52  thus, unfolding BCX using single molecule AFM represents the first study in which the  elucidation of folding kinetics is possible. The folding behaviour of BCX revealed in this manner demonstrates that folding of BCX may be described using a single rate constant. However, the unfolding of BCX has been previously characterized according to the pH dependent Tm observed by circular dichroism or calorimetry. This thermal stability is highly variable according to the different types of xylanases studied.53 The xylanase family comprises a large number of proteins, a number of which exhibit resistance to environmental conditions typically unfavorable to the majority of xylanase members. Elucidating the unfolding and folding kinetics of BCX could lead to a similar comparative study between BCX and its extremophilic cousins, examining how adaptations that allow other proteins to flourish within adverse conditions affect the manner in which folding or unfolding occurs. 3.3.3 Determining the Mechanical Effects of Inhibitor Binding A number of inhibitors have previously been found that bind BCX in a manner that mimics the covalent glycosyl-enzyme intermediate formed as BCX binds xylan. The binding of these inhibitors is long-lived, and has allowed for the elucidation of the detailed mechanism of BCX, including the specific structural interactions responsible for the enzymatic functionality of BCX. In order to investigate how the mechanical properties of BCX are affected by substrate binding, the inhibitor 2,4-dinitrophenyl-4'-amino-2-F-xylobioside was applied in excess to (GB1)4BCX(GB1)4 adsorbed onto glass in PBS (pH~6). The inhibitor, which was graciously provided by Stephen G. Withers, was applied in concentrations in excess of those used in previous NMR studies, and the duration of experiments shortened to less than six hours in accordance with the half life of the bound inhibitor (t1/2~330 min).41 Unfolding of the heteropolyprotein was done using single pull experiments, where a single molecule of polyprotein is picked up between the surface and tip, extended until detachment occurs and the experiment repeated with a new single molecule of protein. BCX peaks were identified as low extension unfolding events, and compiled according to their resultant ∆Lc and average unfolding force (Figure 3.9).  106  The force histogram of compiled unfolding events from BCX bound to 2,4-dinitrophenyl4'-amino-2-F-xylobioside demonstrated an average unfolding force of 58 ± 31 pN (n=61), comparable to that of unbound BCX. Compiling the ∆Lc resulting from fitting BCX unfolding events using the Worm-Like chain resulted in an apparent bi-modal distribution. Within this, most unfolding events occurred exhibiting a ∆Lc of 65 ± 6 nm, comparable to that of unbound BCX. A smaller population of events could be fit to an average ∆Lc of 52 ± 6 nm. This effect may be amplified by the relatively low number of unfolding events compared for inhibitor bound BCX as compared to that of the unbound form, where further experimentation may further resolve the two ∆Lc populations.  107  Figure 3.9 Unfolding of Inhibitor Bound BCX A Inhibitor  B 10  No. of Events  53 nm  18 nm  8 6 4 2 0 0  20  40  60  80  100  120  140  Force (pN)  C 10  No. of Events  64 nm  18 nm  8 6 4 2  200 pN 50 nm  0 30  40  50  60  70  80  90  ∆Lc (nm)  Figure 3.9 The nanomechanical behaviour of BCX bound to inhibitor. The inhibitor 2,4-dinitrophenyl-4'amino-2-F-xylobioside was applied to BCX, which was subsequently unfolded using single molecule AFM. The force extension curves of [GB1]4BCX[GB1]4 bound to inhibitor reveal unfolding events that occur at lower ∆Lc (top three traces, Figure 3.9 A), as well as events that occur with the typical ∆Lc of BCX (bottom two traces, Figure 3.9 A). This is reflected in the histogram of ∆Lc, which demonstrates two populations of ∆Lc (52 ± 6 nm, black fit in Figure 3.9 C; 65 ± 6 nm, red fit in Figure 3.9 C). The average unfolding force of inhibitor-bound BCX (58 ± 31 pN, Figure 3.9 B) was not significantly different from that of unbound BCX.  3.3.3.1 Examining the Effects of Inhibitor Binding on Unfolding Force As observed, the binding of the transition state inhibitor 2,4-dinitrophenyl-4'-amino-2-Fxylobioside did not drastically alter the unfolding force observed as bound BCX was forcibly unfolded. However, unfolding of inhibitor bound BCX did reveal the presence of a smaller population of unfolding events that unfolded at a lower value of ∆Lc than that of the unbound enzyme. Previous investigations of the enzyme mouse dihydrofolate reductase bound to a catalytic inhibitor in a similar manner revealed the increased population of certain unfolding 108  transitional states. The bound enzyme was unfolded using single molecule AFM without significant change in the protein’s mechanical stability.54 That inhibitor binding does not significantly affect the observed mechanical stability of a protein can be attributed to the equal stabilization of the native state of the protein relative to that of the unfolded state. This results in no net mechanical stabilization, and no change in the observable mechanical stability of the bound protein. 54 The presence of the smaller distribution of BCX unfolding events that occur with a ∆Lc~52 nm may show the prevalence of this corresponding unfolding intermediate state as inhibitor bound BCX unfolds in a similar manner as to that of mouse dihydrofolate reductase. It must be noted that the number of bound-BCX unfolding events examined was much smaller than the number scrutinized in the case of unbound BCX. Thus, further study of the behaviour of the bound BCX protein is warranted before such a mechanism may be unequivocally assigned. 3.3.4 Investigating the Effects of Structural Changes on the Mechanical Behaviour of BCX Three mutants of BCX were constructed and engineered into a heteropolyprotein as was done for the wild type BCX, creating three individual (GB1)4BCXMutant(GB1)4 constructs. Investigating these mutant constructs was twofold: firstly, investigate how the mechanical stability of BCX is affected by the insertion of covalent bonds within its structure, and how increased bonding affects the apparent variability observed within the contour length increment of wild type BCX. The three disulfide mutants, (GB1)4BCX10x36(GB1)4, (GB1)4BCX61x177(GB1)4and (GB1)4BCX100x148(GB1)4, were chosen according to proximal locations for cysteine residue insertion such that disulfide bonds would form spontaneously. One of these mutants, BCX100x148, has previously been shown to increase the thermal stability of BCX,43 presenting an ideal model system for determining how this same mutation affects the mechanical properties of BCX. All three of the mutant BCX constructs was investigated in the same manner as the wild-type construct using single molecule AFM. However, due to the low pickup affinity of these proteins, refolding was more difficult than that of the wild type construct, subsequently necessitating the use of single pull experiments. Low pickup also made it difficult to collect sufficient unfolding events to create reliable statistics; the majority of experimental effort thus centered upon the (GB1)4BCX100X148(GB1)4 construct previously demonstrated to increase the thermal stability in comparison with wild type BCX.43  109  Figure 3.10 Investigating the Force Extension Fingerprint of BCX100x148  A  64 nm  B  64 nm  18 nm  C  53 nm  18 nm  200 pN 50 nm  Figure 3.10 The nanomechanical behaviour of the disulfide bonded mutant BCX100x148. This disulfide bonded mutant was constructed in a similar manner as for that of wild type BCX, in which the BCX component of the polyprotein is book ended by four repeats of GB1 (Figure 3.10 A). A disulfide bond was engineered between residue 100 and 148 along the primary structure of BCX by introducing two site mutations: S100C and N148C (Figure 3.10 B). Introduction of disulfide bonds into the structure of a protein should decrease the resultant ∆Lc as covalent bonds are too strong to be affected by force generated by AFM. This effect is noted within the force extension curve of the disulfide bonded mutant (Figure 3.10 C bottom traces, green fit lines). However, unfolding events occurring at the ∆Lc of wild type BCX were identifiable, demonstrating the incomplete nature of disulfide bond formation.  Unfolding events of BCX100x148 were identified from the force extension of (GB1)4BCX100x148(GB1)4, occurring at low extension as that of wild type BCX. These unfolding events were fit using the Worm-Like chain model of polymer elasticity; compiling the values obtained for ∆Lc into a histogram results in two populations of ∆Lc. The larger population may be fit to a ∆Lc of 52 ± 9 nm, corresponding fairly well with the ‘trapping’ of 48 amino acids by the formation of the covalent disulfide bond between cysteine residues engineered at S100C and N148C. This population exhibiting a decreased value of ∆Lc suggests that disulfide bonds 110  were formed within the majority of BCX molecules unfolded. A small population of BCX molecules examined were observed to unfold with a ∆Lc of 63 ± 4 nm, representing the definable minority of BCX100X148 mutants that did not form a bridging disulfide bond. Compiling the unfolding events of BCX100X148 resulted in a force histogram where unfolding forces were centered about 56 ± 30 pN (n=130), comparable with that of wild type BCX. Figure 3.11 The Average Unfolding Force and ∆Lc of BCX100x148 B  A 14  12  No. of Events  No. of Events  12 10 8 6 4  10 8 6 4 2  2  0  0 0  20  40  60  80  Force (pN)  100  120  140  20  30  40  50  60  70  80  90  ∆Lc (nm)  Fi gure 3.11 The nanomechanical behaviour of the disulfide mutant BCX100x148. Introduction of a disulfide bond at position 100 and 148 along the primary structure of BCX did not affect the average unfolding force observed (Figure 3.11 A). The average unfolding force of the 100x148 BCX disulfide mutant was 56 ± 30 pN (n=130). The contour length increment of this protein reveals two populations (Figure 3.11 B): a larger population at 52 ± 9 nm and a smaller one at 63 ± 4 nm. The population of molecules exhibiting a reduced ∆Lc correspond to that expected with the introduction of a covalent disulfide bond at the 100x148 position, where the smaller population corresponds to molecules that did not form this bridging disulfide bond.  3.3.4.1 The Mechanical Effects of Introduced Bonding: Engineered Disulfide Bonds Engineering structural disulfide bonds through the introduction of proximal cysteine residues in the primary structure has previously been observed to increase the thermal and mechanical stability of a variety of proteins, including the thermal stability of BCX.43 The introduction of a disulfide bond between positions 100-148 in the structure of BCX not only increased its thermal stability, but also allowed the mutant enzyme to refold after the application of heat in a way that allows the refolded enzyme regain its functionality.43 This mutant was thus an ideal system in which to examine how this same mutation would affect mechanical properties compared with that of wild type BCX. The average unfolding force of BCX100X148 (~56 pN) 111  did not deviate from that exhibited by wild type BCX (~55 pN). Thus, it appears as though the mechanical stability of BCX is unaffected by the insertion of a covalent bond within this particular location in its structure. The ∆Lc exhibited by this mutant demonstrated the presence of this covalent bond with a large population of unfolding events fit using a lower contour length increment than that of wild type of BCX. No other significant populations of ∆Lc were noted within the unfolding data compiled for this protein, signifying a lack of noteworthy intermediate states forming as BCX100x148 unfolds. The lack of effect noted for the mechanical stability of this mutant seems also to extend to the contour length increment demonstrated as the protein unfolds; thus, the overall mechanical behaviour of BCX is seemingly unchanged by the insertion of a covalent bond between position 100 and 148 along its primary structure. 3.4 Conclusion Using single molecule AFM, the mechanical properties of the non-mechanical protein BCX have been elucidated as outlined. This demonstrates that BCX exhibits a low unfolding force and variable contour length increment in much the same manner as other non-mechanical proteins previously examined using single molecule AFM. As AFM experiments require small concentrations of protein, and examine one molecule at a time, aggregation effects previously observed within ensemble measurements have been eliminated, allowing for the exposition of the unfolding and folding kinetics of BCX. Illuminating these properties allows for the opportunity to compare between the properties of different non-mechanical proteins in an attempt to reveal how mechanical stability is encoded at a molecular level. It also allows for the possibility of further studies of other members of the xylanase family exhibiting extremophilic behaviour to clarify how these attributes affect exhibited mechanical stability. The mechanical properties of BCX bound to the 2,4-dinitrophenyl-4'-amino-2-F-xylobioside inhibitor were also explicated, revealing the possible population of unfolding intermediate states as inhibitor bound BCX is unfolded under force. Finally, the effects of inserting a disulfide bond in a location known to affect the thermal stability of BCX was examined, where it was determined that such a mutation had little to no effect on the relative unfolding force when compared to wild type BCX.  112  3.5 Materials and Methods 3.5.1 Construction of BCX-Containing Proteins 3.5.1.1 Construction of the (GB1)4BCX(GB1)4 Plasmid The heteropolyprotein construct (GB1)4BCX(GB1)4 was engineered in a similar manner as was described for T4 lysozyme,46 utilizing the same molecular biology protocol as previously described for tenascin-X. A plasmid containing BCX was generously given by Laurence McIntosh; this copy of BCX ascribed a semi-synthetic sequence that had been optimized for expression yet exhibited the same mechanistic and structural properties as ‘natural’ BCX. This plasmid was amplified and engineered to contain 5’ BamHI, and 3’ KPNI and BglII restriction sites using a polymerase chain reaction (PCR) that utilized a 5’ primer of sequence CGC GGA TCC GCT AGC ACA GAC TAC TGG C AAC GTT GGA AG.  and 3’ primer of sequence CGC GGT ACC GCA ACA AGA TCT CCA CAC TGT  Subsequent PCR products were sequenced using direct DNA sequencing to  confirm successful amplification. Constructing the final nonameric BCX-containing gene product involved three alternating digestion and ligation steps in a manner similar to that described for tenascin-X. The BCX gene containing the requisite restriction sites was isolated by digesting PCR amplification products using the restriction enzymes BamHI and KNPI. Digestion in this manner resulted in the creation of overhanging ‘sticky ends’ whose sequence corresponds to that of the vector pUC19(GB1)4 digested with BglII and KPNI. The sticky ended BCX insert was subsequently ligated into the digested pUC19(GB1)4 vector using T4 DNA ligase to form pUC19(GB1)4BCX. This plasmid was subsequently cut by BamHI and KPNI digestion to create the sticky-ended insert (GB1)4BCX, which was subsequently ligated into the linearized expression vector pQE80L. The original pUC19(GB1)4 plasmid was cut using BamHI and KNPI restriction sites to form the (GB1)4 gene insert and the vector pQE80L(GB1)4BCX made via digestion with KNPI and BglII. The final plasmid construct pQE80L(GB1)4BCX(GB1)4 was constructed by ligating the sticky ends of the (GB1)4 insert with the pQE80L(GB1)4BCX vector. The correct size of the constructed gene was verified using agarose gel electrophoresis, in which the linear 113  gene containing (GB1)4BCX(GB1)4 was made by digesting the engineered pQE80L(GB1)4BCX(GB1)4 vector with BamHI and KPNI restriction enzymes, and running a portion of the digested DNA insert on a 1% agarose gel. The size of the insert was compared to that of a pre-stained DNA ladder also run on the gel. 3.5.1.1.1 Protein Expression of (GB1)4BCX(GB1)4 Expression of (GB1)4BCX(GB1)4 was done by over-expression within a DH5α strain of E.Coli. Approximately 200 µL of the soluble expression vector pQE80L(GB1)4BCX(GB1)4 containing the gene to be expressed was combined with a similar amount of DH5α cells on ice. Exposing the bacterial cells to heat results in the opening their nuclear pores, which allows plasmid DNA to be inserted into the nucleus of the competent cells. Expression of the desired gene involves utilization of the machinery within the bacterial hosts to make the final protein product from the engineered gene. Thus, transcription and translation of the engineered BCX plasmid into protein is accomplished by optimizing the growing conditions of bacteria that contain the desired gene. Also contained within the engineered BCX plasmid is a gene encoding for antibiotic resistance; bacteria cultures were thus grown in 37̊C nutrient broth solutions containing 1M ampicillin. Addition of antibiotic to the nutrient broth ensures that only bacteria containing the plasmid encoding for antibiotic resistance, and thus the gene of interest, can grow within the nutrient broth. Bacterial cultures were incubated until an optical density of approximately 0.6 was attained (λ=600 nm, approximately 3 hours). At this time, expression of the BCX containing gene was induced by the addition of 1M IPTG, which binds to upstream repressor elements on the BCX plasmid. Induced bacterial cultures were incubated for an additional four hours at 37̊C. During the incubation period, bacteria containing the BCX plasmid replicated and utilized the machinery present within the bacterial host to transcribe and translate the engineered BCX-containing gene into a BCX-containing protein. Subsequent steps focused on the isolation of this protein construct. Host bacterial cells were lysed within a 1M solution of lysozyme. Lysed cells were then treated with DNase and RNase solutions, which digested DNA and RNA fragments, and subsequently centrifuged to separate insoluble elements from the bacterial cell, such as the cellular membrane, from the soluble protein located in the supernatant. The soluble 114  protein was purified from the cell lysis supernatant by passing the supernatant through a Co2+ affinity column. Binding of the protein of interest was accomplished through the affinity between the metal resin and the six histidine repeats located on the N-terminus of the engineered protein. Elution of the bound protein was accomplished by adding an excess of a PBS solution containing imidazole. The eluted protein was stored at 4˚C in a phosphate buffered saline solution at a concentration of 1.0 µg/ml. 3.5.1.2 Construction of Disulfide Bonded Mutants Three disulfide mutants of BCX were engineered into a similar polyprotein chimera as described for wild type BCX. This was done in a similar manner as for the wild type, save for initial steps that utilized a ‘megaprimer’ approach to insert cysteine residues into the primary sequence of BCX.55 The position of residues likely to result in the formation of a disulfide bond was identified using the DSDBase disulfide database (http://caps.ncbs.res.in/dsdbase//modip.html). Potential sites of the insertion of cysteine were identified on the basis of their proximity and how likely the subsequent formation of a disulfide bond was. Six sites were identified as likely to form three separate disulfide bonds: T10 and F36, N61 and S177, and S100 and N148. The megaprimer approach entails an extra round of PCR in order to engineer two different site mutations into each of the three desired disulfide mutants. Two primers were made for each desired protein (Table 3.1), each primer containing the point mutation necessary such that the triplet base pair at the specific residue location encodes for cysteine rather than the wild type residue. The first round of PCR utilized BCX with attached 5’ BamHI and 3’ KPNI and BglII sites as a template and both types of primer; the product of this first round of PCR is short linear dSDNA containing both of the mutations necessary to form each of the three disulfide bonds. Template DNA was digested using dPN1 after the first PCR reaction was completed. The second PCR stage used mutant dSDNA as primers and the same template BCX as the initial PCR stage to create copies of the BCX gene containing both mutations, and the PCR products transformed into XLI Blue competent cells. The presence of both cysteines within the sequence of all three plasmids was ascertained by direct DNA sequencing. The subsequent engineering of gene constructs and the final expression of three disulfide-containing mutant protein chimeras was done as described for wild type BCX. Final protein concentrations ranged dependent upon the specific construct: 115  (GB1)4BCX10X36(GB1)4 showed a final concentration of 0.5 µg/mL, (GB1)4BCX100x148(GB1)4 at 0.9 µg/mL, and (GB1)4BCX61x177(GB1)4 at 0.6 µg/mL. Table 3.1 Primers Used in the Construction of Disulfide Mutants Disulfide Primer Type  Position  Primer Sequence  T10C Forward CAAAATTGGTGTGATGGGGGAGGTATAG F36C  Reverse  GTCTAATACCGGAAATTGTGTTGTTG  N61C Forward  GCCGTGTGGCAATGGATATTTAAC  S177C Reverse  GACCGAAGGCTACCAGTGCTCTG  S100C Forward  GTAAAATGTGATGGGGGTACATATG  N148C Reverse  CACCATTACGTTCACCTGTCACGTG  3.5.2 AFM Experiments All AFM experiments were conducted in a similar manner as described previously in this document. Briefly, approximately 1 µL of protein in solution was applied to cleaned glass surface and allowed to adsorb for ~5 minutes. After this time, excess PBS was pipetted from the glass surface and 50 µl of fresh PBS (pH~6) applied to the surface. Pipetting excess PBS from the glass surface acts to reduce the concentration of protein that is free in solution. Calibration of the silicon nitride cantilevers typically resulted in a spring constant of ~50 pN/nm. 3.5.3 Inhibitor Binding to BCX for AFM Experiments Approximately 0.6 mg of 2,4-dinitrophenyl-4'-amino-2-F-xylobioside inhibitor was obtained through generous collaboration with Steven G. Withers; this white crystalline powder was subsequently dissolved in 1 mL of ddH20 and stored at -80˚C until used in AFM experiments. The manner used to bind inhibitor to BCX for AFM experiments was adapted from that reported from previous studies using bound forms of BCX.40 For experiments involving the use of inhibitor, 1 µL of BCX solution was added to a clean glass cover slip and 116  allowed to adsorb for approximately five minutes. After this time, 5 µL (3 µg) of inhibitor solution was added and allowed to bind to completion for approximately 15 minutes as indicated by previous NMR binding studies.40 Inhibitor binding to BCX solution has previously been noted to result in a yellow solution colour that was not observed, possibly due to the low concentration of both BCX and its inhibitor as well as the grey colour of the AFM stage, which effectively obscures the ability to note any solution colour change. After 15 minutes, AFM experiments were carried on as previously described.  117  3.6 References 1. Prade, R. A., Xylanases: from biology to biotechnology. Biotechnol Genet Eng Rev 1996, 13, 101-31. 2. 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G., The pKa of the general acid/base carboxyl group of a glycosidase cycles during catalysis: a 13C-NMR study of bacillus circulans xylanase. Biochemistry 1996, 35, (31), 9958-66. 41. Miao, S.; Ziser, L.; Aebersold, R.; Withers, S. G., Identification of glutamic acid 78 as the active site nucleophile in Bacillus subtilis xylanase using electrospray tandem mass spectrometry. Biochemistry 1994, 33, (23), 7027-32. 42. Davoodi, J.; Wakarchuk, W. W.; Campbell, R. L.; Carey, P. R.; Surewicz, W. K., Abnormally high pKa of an active-site glutamic acid residue in Bacillus circulans xylanase. The role of electrostatic interactions. Eur J Biochem 1995, 232, (3), 839-43. 43. Wakarchuk, W. W.; Sung, W. L.; Campbell, R. L.; Cunningham, A.; Watson, D. C.; Yaguchi, M., Thermostabilization of the Bacillus circulans xylanase by the introduction of disulfide bonds. Protein Eng 1994, 7, (11), 1379-86. 44. Connelly, G. P.; Withers, S. G.; McIntosh, L. P., Analysis of the dynamic properties of Bacillus circulans xylanase upon formation of a covalent glycosyl-enzyme intermediate. Protein Science 2000, 9, (3), 512-524. 45. Junker, J. P.; Hell, K.; Schlierf, M.; Neupert, W.; Rief, M., Influence of substrate binding on the mechanical stability of mouse dihydrofolate reductase. Biophysical Journal 2005, 89, (5), L46-L48. 46. Peng, Q.; Li, H., Atomic force microscopy reveals parallel mechanical unfolding pathways of T4 lysozyme: evidence for a kinetic partitioning mechanism. Proc atl Acad Sci U S A 2008, 105, (6), 1885-90. 47. Weaver, L. H.; Matthews, B. W., Structure of bacteriophage T4 lysozyme refined at 1.7 A resolution. J Mol Biol 1987, 193, (1), 189-99. 48. Best, R. B.; Li, B.; Steward, A.; Daggett, V.; Clarke, J., Can non-mechanical proteins withstand force? Stretching barnase by atomic force microscopy and molecular dynamics simulation. Biophys J 2001, 81, (4), 2344-56. 49. Oberhauser, A. F.; Carrion-Vazquez, M., Mechanical biochemistry of proteins one molecule at a time. J Biol Chem 2008, 283, (11), 6617-21. 50. Brockwell, D. J.; Paci, E.; Zinober, R. C.; Beddard, G. S.; Olmsted, P. D.; Smith, D. A.; Perham, R. N.; Radford, S. E., Pulling geometry defines the mechanical resistance of a betasheet protein. at Struct Biol 2003, 10, (9), 731-7. 51. Carrion-Vazquez, M.; Li, H.; Lu, H.; Marszalek, P. E.; Oberhauser, A. F.; Fernandez, J. M., The mechanical stability of ubiquitin is linkage dependent. at Struct Biol 2003, 10, (9), 738-43. 120  52. Nath, D.; Rao, M., Artificial chaperone mediated refolding of xylanase from an alkalophilic thermophilic Bacillus sp. Implications for in vitro protein renaturation via a folding intermediate. Eur J Biochem 2001, 268, (20), 5471-8. 53. Poon, D. K.; Webster, P.; Withers, S. G.; McIntosh, L. P., Characterizing the pHdependent stability and catalytic mechanism of the family 11 xylanase from the alkalophilic Bacillus agaradhaerens. Carbohydr Res 2003, 338, (5), 415-21. 54. Junker, J. P.; Hell, K.; Schlierf, M.; Neupert, W.; Rief, M., Influence of substrate binding on the mechanical stability of mouse dihydrofolate reductase. Biophys J 2005, 89, (5), L46-8. 55. Sharma, D.; Cao, Y.; Li, H., Engineering proteins with novel mechanical properties by recombination of protein fragments. Angew Chem Int Ed Engl 2006, 45, (34), 5633-8.  121  Chapter 4 Comparing the anomechanical Properties of Mechanical and onMechanical Proteins: Elucidating the Molecular Determinants of Mechanical Attributes 4.1 Investigating on-Mechanical and Mechanical Proteins Using Single Molecule AFM as a Means of Illuminating Origins of Mechanical Behaviour Proteins are critical to life, acting as physiological workhorses for an enormous array of biological processes that perform the basic functions necessary for biological life. Comparing the intrinsic complexity and diverse engineering that proteins may exhibit with man-made technology exemplifies their simplistic, ingenious natural design. This simplicity belies the complex factors that contribute to the overall protein blueprint, a delicate balancing act which the scientific community is only beginning to understand. The motivation behind trying to understand and harness protein machinery by understanding the mechanisms by which they fold, as well as how their specific properties and functionality are encoded within their structure, is the desire to make use of these specific functional characteristics developed through millions of years of evolution. Understanding these processes have far ranging consequences, including grasping the basis of diseases involving protein misfolding and the rational design of proteins with novel functional folds. The development of Top7, a protein designed in silico to fold with a completely novel topology, is a step in this direction, where proteins could be rationally designed to act as pharmaceutical agents and novel nanoscale machines.1 Top7 is a step towards rationally tuning the fold and properties of proteins, demonstrating the possibilities of protein engineering but also illustrating the current impediments within this promising field. How proteins fold, and how their properties are delineated within their molecular structure, is still a poorly understood process; easily and rationally tuning these characteristics in a functional manner is still a future goal in the field of protein nanomechanics. Comparative studies between differing functional and structural classes of proteins seek to elucidate details on how information for how folding and functional characteristics are engineered at a molecular level. In the study denoted here, the nanomechanical properties of 122  two proteins, tenascin-X and bacillus circulans xylanase (BCX), were elucidated using single molecule AFM. These proteins are divergent in terms of their molecular functionality, with tenascin-X acting as a mechanical protein intrinsic to the structural functionality of the extracellular matrix and BCX exhibiting non-mechanical enzymatic activity in the cleavage of xylan chains. Briefly examining the nanomechanical properties of both these proteins reveals little similarity. The mechanical stability of tenascin-X, whose unfolding properties originates within its constituent FnIII domains, is greater than that of BCX, signified by an increase in average unfolding force of about 100 pN. This is likewise for unfolding and folding kinetics elucidated previously, with both proteins exhibiting differing methods by which they unfold and fold. The simplest comparison between the two proteins is made between the average unfolding forces exhibited by unfolding events within the force extension curves of each. As the average unfolding force corresponds to the mechanical stability intrinsic to the protein, it is easily noted that tenascin-X is more mechanically stable than BCX. Previous studies have noted a correlation between mechanical stability and structural content of proteins. 2 This is a lesser effect in explaining the comparatively low mechanical stability of BCX observed. Instead, topological effects can aptly describe why tenascin-X unfolds at much higher force than that of BCX. Although the structure of bovine tenascin-X FnIII domains are not known at this time, the high structural homology of FnIII domains means that bovine FnIII domains likely demonstrate a β-barrel structure similar to that of other structures available for FnIII domains. This structure, with its parallel N and C termini, is a classic example of a topology which, when made to unfold, does so in a ‘shearing’ motion correlated with high mechanical stability.3, 4 The topology of BCX aligns its N and C termini in close proximity with one another, making it likely that BCX unfolds with a ‘zippering’ mechanism that classically results in low apparent mechanical stability. Topological effects most likely play a large role in determining the mechanical stability exhibited by an unfolding protein, further exemplified by the number of high stability folds that exhibit structures with parallel terminus likely to undergo ‘shear’ unfolding (Table 1.1). Proteins that are designed to withstand mechanical force are likely engineered such that applications of force are made across multiple bonds within its structure in order to increase their mechanical resilience. This is also an important  123  consideration in the rational engineering of proteins, in that how force is felt through the network of bonds within the structure of a protein appears to be a large determinant for mechanical stability. That the mechanical protein tenascin-X exhibits a higher average unfolding force than the non-mechanical BCX is not surprising considering their physiological roles: if tenascin-X was not able to withstand mechanical force, the structural organization and resiliency of many physiological tissues would be negatively affected. A hint of this is observed within the phenotypic expression of the Ehlers-Danlos syndrome, where patient suffering from the recessive form involving point mutations or deletions within the tenascin-X gene exhibits signs of joint and tissue hyper elasticity, skin that is easily bruised and wounds that heal poorly. It is likely that the mechanical stability observed within tenascin-X is the result of evolutionary pressure, in that tenascin-X whose mechanical stability is low would not be functional within an environment under constant mechanical pressure such as the ECM. These pressures would not be at play for proteins such as BCX, whose function is far removed from any need to exhibit high mechanical stability. In a parallel type of argument, it has been demonstrated that proteins exhibit only marginal values of thermodynamic stability if such an increase in stability did not confer a specific evolutionary advantage, such as resilience towards destabilizing mutations.5-7 Although thermodynamic stability is a poor predictor of mechanical stability, a similar mechanism may also be in play in terms of mechanical stability. Proteins such as BCX have no obvious need to be mechanically stable, exhibiting functionality seemingly independent of transduced or applied mechanical force. This is not to say that non-mechanical proteins cannot exhibit high mechanical stability, only that mechanical stress is not intrinsic to function in the same manner as mechanical proteins such as the tenascins or titin. Nonmechanical proteins that do exhibit high mechanical stability, such as GB1, may do so if mechanical force acts in a more subtle manner within their environment or functionality. High mechanical stability within these proteins may also be incidental, where structural organization into a mechanically stable fold serves another intrinsic function of the protein in a manner that is not easily disconcernable.  124  4.2 Future Studies of Protein anomechanics The future work around attempting to illuminate questions of protein folding and functionality will benefit from the increasing knowledge of the nanomechanical properties of diverse proteins such as BCX and tenascin-X as well as how specific mutations and changes in structure affects these characteristics. To this end, much work within the Li lab is focused on the nanomechanical effects of structural changes within proteins previously observed using single molecule AFM. The use of in silico modeling techniques is the perfect counterpoint for such studies. Techniques such as molecular dynamics simulation are able to provide the molecular explanation behind the experimental properties observable using AFM. Computer simulation of protein unfolding, paired with experimental data gained from AFM provides the possibility of providing a detailed molecular explanation behind why certain mechanical behaviors are observed. The power of these combined techniques has been previously demonstrated in studies such as the prediction of key H-bonds to the mechanical stability of FnIII10 domains from fibronectin.8 Future work towards simulating the unfolding of tenascinX FnIII domains as well as BCX could help establish whether unfolding topology is indeed the critical factor in delineating their drastically different mechanical stabilities. Advances in the computational power available will continue to increase the efficiency and strength of such simulations, making them increasingly valuable in the field of protein nanomechanics. Elucidating the nanomechanical properties of tenascin-X and BCX could have implications other than how these proteins encode their mechanical properties. Studying the behaviour of wild type tenascin-X allows for the future study of how disease causing mutation affects mechanical stability observable using AFM. This could be accomplished by examining how point mutations known to cause the recessive form of Ehlers-Danlos Syndrome affects mechanical stability in comparison to that exhibited by the wild type protein. Also of future interest is the further elucidation of mechanical properties of FnIII domains from multiple protein sources, and how domains with high structural but low sequence homology encode differing mechanical functionality. Comparing differing species of tenascin, such as was done for tenascin-C and tenascin-X, could also further develop how these proteins encode for 125  different characteristics beginning within a similar structural blueprint. Studying the mechanical behaviour of BCX demonstrated the advantages present when using AFM, in that a protein whose tendency towards aggregation meant that ensemble folding measurements were impossible was evaluated in terms of its previously unfeasible folding kinetics. The question of how proteins fold, and what gives them their nanomechanical properties, is a difficult fundamental one offering no easy answer. However, the knowledge base of what is understood concerning protein folding and their nanomechanical properties is perceptibly growing fuelled in part by research such as presented here. That the field is inching towards a solution is obvious by the increased sophistication and deeper understanding of the techniques used as well as the information they are able to impart. Illuminating the forces driving proteins to fold and how function is attained may be a thorny proposition, but one that is becoming more and more feasible as the field progresses and that has huge potential to affect the world in which we live.  126  4.3 References 1. Kuhlman, B.; Dantas, G.; Ireton, G. C.; Varani, G.; Stoddard, B. L.; Baker, D., Design of a novel globular protein fold with atomic-level accuracy. Science 2003, 302, (5649), 1364-8. 2. Oberhauser, A. F.; Badilla-Fernandez, C.; Carrion-Vazquez, M.; Fernandez, J. M., The mechanical hierarchies of fibronectin observed with single-molecule AFM. J Mol Biol 2002, 319, (2), 433-47. 3. Brockwell, D. J.; Beddard, G. S.; Paci, E.; West, D. K.; Olmsted, P. D.; Smith, D. A.; Radford, S. E., Mechanically unfolding the small, topologically simple protein L. Biophys J 2005, 89, (1), 506-19. 4. Carrion-Vazquez, M.; Li, H.; Lu, H.; Marszalek, P. E.; Oberhauser, A. F.; Fernandez, J. M., The mechanical stability of ubiquitin is linkage dependent. at Struct Biol 2003, 10, (9), 738-43. 5. Bloom, J. D.; Labthavikul, S. T.; Otey, C. R.; Arnold, F. H., Protein stability promotes evolvability. Proc atl Acad Sci U S A 2006, 103, (15), 5869-74. 6. Bloom, J. D.; Silberg, J. J.; Wilke, C. O.; Drummond, D. A.; Adami, C.; Arnold, F. H., Thermodynamic prediction of protein neutrality. Proc atl Acad Sci U S A 2005, 102, (3), 60611. 7. Nikolova, P. V.; Wong, K. B.; DeDecker, B.; Henckel, J.; Fersht, A. R., Mechanism of rescue of common p53 cancer mutations by second-site suppressor mutations. EMBO J 2000, 19, (3), 370-8. 8. Craig, D.; Gao, M.; Schulten, K.; Vogel, V., Tuning the mechanical stability of fibronectin type III modules through sequence variations. Structure 2004, 12, (1), 21-30.  127  

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