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Length distribution of myosin filaments in smooth muscle and implications in the structure and function… Liu, Jeffrey Chao-Yu 2015

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   Length distribution of myosin filaments in smooth muscle and implications  in the structure and function of contractile units  by Jeffrey Chao-Yu Liu B.Sc., The University of British Columbia, 2012  A THESIS SUBMITTED IN PARTIAL FULLFILLMENT OF  THE REQUIREMENT FOR THE DEGREE OF   MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POST DOCTORAL STUDIES (Experimental Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)    February 2015  ©  Jeffrey Chao-Yu Liu, 2015ii  ABSTRACT  Smooth muscle is an essential component of the walls of numerous hollow or tubular organs throughout the body, including blood vessels, airways, and the bladder.  Proper physiological functioning of these organs relies heavily on the appropriate activation and contraction of the smooth muscle tissue.  Pathophysiological conditions may arise from both excessive and insufficient smooth muscle contraction.   Muscle function is closely associated with muscle structure.  More specifically, during a contraction, cyclic interactions between myosin cross-bridges and actin filaments allow for muscle shortening and force generation.  Myosin molecules from smooth muscle and non-muscle cells are known to self-assemble into side-polar filaments in vitro.  However the in situ mechanism of filament assembly is not clear and the question of whether there is a unique length for myosin filaments in smooth muscle is still under debate.  In this study we measured the lengths of 16,587 myosin filaments in three types of smooth muscle cells using serial electron microscopy (EM).  Sheep airway and pulmonary arterial smooth muscle as well as rabbit carotid arterial smooth muscle were fixed for EM and serial ultra-thin (50-60 nm) sections were obtained.  Myosin filaments were traced in consecutive sections to determine their lengths.  The results indicate that there is not a single length for the myosin filaments; instead there is a wide variation in lengths. The plots of observation frequency versus myosin filament length follow an exponential decay pattern.   The most significant finding of this study is that myosin filaments in smooth muscle do not have a uniform length and analysis suggests that the distribution of filament length is iii  a result of a dynamic equilibrium between polymerization and de-polymerization of myosin molecules driven by predictable probabilities of the myosin dimers to bind with and dissociate from each other. iv  PREFACE  This thesis is based on experiments performed by myself in the laboratory of Dr. Paré and Dr. Seow in the Centre for Heart Innovation at St. Paul’s Hospital. The work presented in this thesis is based on published research: Liu, Jeffrey C.-Y., Jörg Rottler, Lu Wang, Jenny Zhang, Chris D. Pascoe, Bo Lan, Brandon A. Norris, Ana M. Herrera, Peter D. Paré, and Chun Y. Seow. 2013. “Myosin Filaments in Smooth Muscle Cells Do Not Have a Constant Length.” The Journal of Physiology 591 (Pt 23): 5867–78.   Reprinted with appropriate permission of the journal.  The experiments were designed and modified by Dr. Seow and me.  Muscle fixation and electron microscopy were performed by Jenny Zhang.  Data acquisition was performed by me and analysis with help from Dr Seow.  The mathematical model explaining the data was developed by Dr Seow and Dr Rottler.  The published manuscript was written by Dr. Seow and me, with contribution from all co-authors. The work presented in this thesis is covered under the Human Ethics Board of UBC certificate number H12-02487, project title “Plasticity in airway smooth muscle” v  TABLE OF CONTENTS   Abstract ................................................................................................................................ ii Preface................................................................................................................................. iv Table of Contents ................................................................................................................. v List of Figures .................................................................................................................... vii List of Abbreviations .......................................................................................................... ix Acknowledgments................................................................................................................ x  Dedication ........................................................................................................................... xi  CHAPTER 1 :  Smooth muscle structure and physiology .............................................1 1.1 General introduction ..........................................................................................1  1.1.1 Smooth muscle ....................................................................................1  1.1.2 General structure of smooth muscle cells ...........................................2 1.2 Ultrastructure of smooth muscle cells................................................................3  1.2.1 Cell membrane ....................................................................................3  1.2.2 Organelles ...........................................................................................7  1.2.3 Cytoskeleton......................................................................................10  1.2.3.1 Organization of filaments...................................................10  1.2.3.2 Dense bodies and dense plaques ........................................15  1.2.3.3 Contractile apparatus – smooth muscle “sarcomere  ..........16 1.3 Smooth muscle contraction ..............................................................................20 1.4 Plasticity and length adaptation .......................................................................23  CHAPTER 2:  Hypothesis, specific aims and rationale ...............................................25  2.1  Hypothesis.......................................................................................................25  2.2  Specific aims ...................................................................................................27  CHAPTER 3:  Materials and methods ..........................................................................29  3.1  Tissue sample ..................................................................................................29 vi   3.2  Tissue preparation ...........................................................................................29  3.3  Electron microscopy .......................................................................................30  3.4  Image analysis.................................................................................................32  3.5  Myosin filament length measurement.............................................................32  3.6  Statistical analysis ...........................................................................................35  CHAPTER 4:  Results .....................................................................................................36  4.1  Control observation.........................................................................................36  4.2  Inter-observer differences ...............................................................................38  4.3  Mathematical model for linear polymerization and fragmentation ................40  4.4  Myosin filaments in relaxed airway smooth muscle.......................................42  4.5  Myosin filaments in relaxed arterial smooth muscle ......................................45 4.6  Filament length measured in longitudinal sections.........................................47  4.7  Myosin filaments in activated smooth muscle ................................................49  CHAPTER 5:  Discussion................................................................................................51  5.1  Discrepancies with previous studies on myosin filament length ....................51 5.2  Proposed model of a smooth muscle contractile unit .....................................53 5.3  Facilitation of myosin filament polymerization in vivo .................................56 5.4  Implications of the mathematical model for linear polymerization ................57 5.5  Limitations of the measurement technique .....................................................59 5.6  New role for myosin in smooth muscle length adaptation .............................60 5.7 Final conclusion ...............................................................................................60  BIBLIOGRAPHY ............................................................................................................61 APPENDIX I ....................................................................................................................67 vii  LIST OF FIGURES  Figure 1:  Membrane specializations on a transverse section of an airway smooth muscle cell ........................................................................................................................................6 Figure 2:  Organelles in a transverse section of an airway smooth muscle cell .................9 Figure 3:  Electron micrograph of transverse section of a tracheal smooth muscle cells  .15 Figure 4:  Schematic illustration of a myosin monomer and a filament ...........................16 Figure 5:  Structural comparison of skeletal and smooth muscle cells .............................18 Figure 6:  A proposed configuration of smooth muscle contractile unit ..........................19 Figure 7: Smooth muscle contractile pathway..................................................................22 Figure 8:  Hypothesized configuration of smooth muscle contractile unit with        myosin filaments of variable lengths .................................................................................26 Figure 9:  Measurement of myosin filament length in serial sections of electron micrographs........................................................................................................................34 Figure 10:  Serial cross sections of mouse diaphragm (skeletal muscle)..........................37 Figure 11:  Inter-observer differences in the measurement of myosin filament lengths  ..39 Figure 12:  Distribution of observation frequency over myosin filament lengths  measured through tracing of consecutive serial cross-sections sheep tracheal           smooth muscle cells ...........................................................................................................44 viii  Figure 13:  Distribution of observation frequency over myosin filament lengths  measured through tracing of consecutive serial cross-sections of arterial smooth      muscle cells ........................................................................................................................46 Figure 14:  Comparison of myosin filament length distributions measured from      tracing serial cross-sections and from longitudinal section with direct length   measurement in tracheal smooth muscle cells ...................................................................48  Figure 15:  Distribution of observation frequency versus myosin filament lengths measured in ACh-activated sheep tracheal smooth muscle ...............................................50 Figure 16:  Schematic illustration of an actin filament lattice containing myosin    filaments with variable lengths ..........................................................................................55          ix  LIST OF ABBREVIATIONS  Ach  - Acetylcholine ASM  - Airway smooth muscle ATP  - Adenosine triphosphate Ca2+  - Calcium EM  - Electron microscopy ER  - Endoplasmic reticulum IP3  - Inositol 1,4,5-triphosphate MgATP - Magnesium adenosine triphosphate MLC20 - Myosin regulatory light chain MLCK  - Myosin light chain kinase MLCP  - Myosin light chain phosphatase OF  - Observation frequency p  - Probability of bonds forming P  - P-value PKC  - Protein kinase C q  - Probability of bonds breaking r  - Mean probability of bond interaction SE  - Standard error SMC  - Smooth muscle cell SR  - Sarcoplasmic reticulum TEM  - Transmission electron microscopy   x  ACKNOWLEDGEMENTS  I owe my thanks to my supervisors, Dr Chun Seow and Dr Peter Paré, for giving the opportunity and privilege to work and study under their guidance.  For two years, they have consistently provided immense support and guidance leading up to the completion of my graduate degree.   I also thank my supervisory committee members, Dr Jordan Guenette and Dr Robert Schellenberg, for their valuable input and assistance, both scientifically and professionally.   I extend my thanks to my fellow Seow lab members.  Particularly, Dr Lu Wang for her patience and advice through the various projects I’ve been involved with during my study.  Also, thanks to Jenny Zhang for her endless hours with fixation and on the electron microscope, just to provide images for me to analyze.  Furthermore, I owe special thanks to Dr Bo Lan, Dr Chris Pascoe, Brandon Norris, and Nick Swyngedouw for their friendship over the years.   Most importantly, I would like to thank my parents, Jack Liu and Alice Wang, and my brother Joseph Liu for their love and support throughout this process.         xi  DEDICATION   I dedicate this work to my family and friends.1  CHAPTER 1:  Smooth muscle structure and physiology  1.1 General introduction 1.1.1 Smooth muscle Smooth muscle is an essential component of various hollow organs such as stomach, urinary bladder, intestines, blood vessels, and airways, playing a major role in various vital functions such as blood pressure regulation and peristalsis.  Smooth muscle is essential in regulating the dynamic shapes and sizes of these hollow organs, therefore dysfunction of smooth muscle can easily lead to various diseases such as systemic hypertension and asthma.  Improved understanding of the physiology of smooth muscle will facilitate our understanding of the pathophysiology of specific diseases associated with smooth muscle dysfunction.    Muscle function is closely associated with muscle structure.  In striated muscle, structure and function are directly correlated due to the presence of highly organized sarcomeres as the basic units in the contractile apparatus.   Smooth muscles, in contrast, lack well-defined and stable contractile units but are still able to generate isometric stress (force per cross-sectional area) comparable to that of skeletal muscle (Gabella 1997).  The absence of organized, repeating structures in smooth muscle makes it difficult to predict changes in mechanical function due to changes in structure.  Another major characteristic that distinguishes smooth from striated muscle is the structural plasticity associated with the former that allows the muscle to produce maximal force over a wide range of lengths via a process called length adaptation (Bossé et al. 2008; Seow 2005a).  This process has 2  been shown to involve the rearrangement of contractile filaments within the cell (Kuo et al. 2003a).  Therefore, the key to understanding the mechanism of this plasticity lies in the identification of the protein structure of the elementary contractile unit within the muscle.  In striated muscle all myosin filaments have the same length.  This allows the basic contractile units in the muscle (sarcomeres) to have the same dimension.  It is not known whether the myosin filaments in smooth muscle have the same length.  Knowing the myosin filament length is an important first step in elucidating the structure of the contractile unit in smooth muscle and is crucial to understanding the relationship between structure and function that underlies the contraction mechanism.  Because the highly organized sarcomeric structure has never been observed in smooth muscle, it is unlikely that myosin filaments in the muscle would have a uniform length.  Rather, it is likely that myosin filaments in smooth muscle would possess a wide range of lengths.  The focus of this thesis research is on determining the length distribution of myosin filaments in vascular and airway smooth muscles, in both relaxed and activated states.       1.1.2 General structure of smooth muscle cells Smooth muscle cells are usually found in relatively organized clusters forming a functional contractile apparatus.  This type of arrangement allows smooth muscle cells to form a mechanical syncytium where proper function is only achievable as a group (Kuo and Seow 2004).  Smooth muscle cells are spindle-shaped and elongated with significantly variable sizes between tissue types and species.  For individual cells (or fibers), dimensions of 100-1000 μm in length and 2-10 μm in diameter at central segment 3  of the cell have been identified (Garfield and Somlyo 1985).  In airway smooth muscle specifically, sizes range from 3-5 μm in diameter and are approximately 250 μm in length (Stephens 2001).   Smooth muscle can be classified based on the type of innervation, single-unit or multi-unit.  In single-unit smooth muscle, an individual nerve innervates multiple cells in a muscle cell bundle with propagation of action potential relying heavily on the presence of gap junctions between neighboring cells.   This type of innervation allows the muscle fibers to contract in unison.  Multi-unit smooth muscle cells, on the other hand, are independent of each other and are required to be innervated individually.  This type of control allows for far more precise and gradual responses.  Airway smooth muscle cells are considered to be intermediate muscles as they are neither purely single-unit nor multi-unit (Stephens 2001).  They have been shown to be richly innervated, much like multi-unit muscle, but still possess considerable amounts of gap junctions for propagation, as in single-unit muscle (Kannan and Daniel 1980).    1.2 Ultrastructure of smooth muscle cells 1.2.1 Cell membrane The smooth muscle cell membrane, or sarcolemma, is an extensible lipid bilayer (~8 nm in thickness) that contains regions of structural specializations that allow for a variety of cellular functions.  The sarcolemma, like all other cell membranes, helps in maintaining structural integrity and facilitates communication with the extracellular environment.  4  Three different types of mutually exclusive structures have been identified on the smooth muscle sarcolemma: caveolae, cell-to-cell junctions, and dense plaques. Caveolae are vesicular invaginations on the surface of the plasma membrane ranging from 50-100 nm in diameter.  Fig. 1 shows several caveolae located along the membrane (black arrowhead). Although their exact function is still unclear, they are believed to aid in signal transduction partly by increasing the surface area of the sarcolemma.  This alteration to the sarcolemma facilitates the exchange of molecules across the membrane (Stephens 2001; Anderson 1998).  Studies have found that several key components of calcium transport, including Ca2+ ATPase and calmodulin, are localized to caveolae (Gabella 1997; Fujimoto 1993).  Furthermore, caveolae are found to be closely associated with the sarcoplasmic reticulum and mitochondria (K. -H. Kuo, Herrera, and Seow 2003b).    These findings suggest that caveolae play an important role in calcium signaling and contraction, much like the T-tubule system in skeletal muscle.   Cell-to-cell junctions are essential connections between adjacent cells that provide both mechanical and electrical coupling.  In smooth muscle, mechanical coupling is provided by adherens junctions made of opposing dense plaques from neighboring cells (also known as intermediate junctions), allowing for effective force transmission between cells (Garfield and Somlyo 1985).  On the other hand, electrical coupling in smooth muscle is provided by gap junctions that permit transmission of excitatory signals between cells.  An example is shown in the inset of Fig. 1.  These junctions are formed by transmembrane pairing of connexon molecules.  Therefore, gap junctions functionally work as molecular channels that facilitate calcium signaling and action potential propagation between smooth muscle cells (Gabella 1997). The abundance of gap 5  junctions varies between smooth muscle types, but has been reported to be 2.7 ± 0.3 (SE) per 100 cells in airway smooth muscle (Stephens 2001). Dense plaques are electron dense regions of the membrane that are closely associated with the cytoskeleton and contractile apparatus (Fig. 1, white arrows).  They will be discussed in detail in a later section.         6    Figure 1:  Membrane specializations on a transverse section of an airway smooth muscle cell.  Areas of the cell membrane occupied by caveolae (black arrow head) and dense plaques (white arrow) appear in an alternating pattern.   Dense bodies are found throughout the cytoplasm (black circle).  A gap junction is marked by two opposing black arrows, where the cells are in very close proximity.  The inset shows a gap junction at a higher magnification.         7  1.2.2 Organelles The nucleus is the largest organelle in smooth muscle, located at the center of the cell as an elongated structure along the longitudinal axis (Fig. 2).  It possesses a double-layer nuclear membrane with pores, which allows for the transport of proteins and RNA between the nucleoplasm and cytoplasm.  In airway smooth muscle, the nuclear membrane has been shown to directly interact with contractile filaments, suggesting potential involvement in force transmission along the longitudinal axis of the cell (Kuo-Hsing Kuo and Seow 2004a).   The endoplasmic reticulum (ER) systems are comprised of a network of sacs and tubules and are situated within the cytoplasm of all smooth muscle cells.  These systems can be categorized into rough or smooth ER depending on whether ribosomes are present or absent.  The ribosome-rich rough ER is the main site for protein synthesis and transport within the cell, while the smooth ER is primarily involved in the synthesis and transport of lipids and steroids.  In smooth muscle, smooth ER is replaced by the sarcoplasmic reticulum (SR), which acts as a compartment for calcium storage as well as a conduit for calcium transport within the cell (K. -H. Kuo, Herrera, and Seow 2003b).  Fig. 2 shows an example of a ribosome-rich rough ER in close proximity to the centrally located nucleus. Mitochondria are large organelles that are relatively abundant in smooth muscle cells, making up approximately 3-10% of the cytoplasmic volume.  They are observed as elongated structures (1-2 μm in length, 0.3 μm in width) and are found in clusters within the cytoplasm with close association with the SR calcium storage sites (Gabella 1997).  8  Fig. 2 shows several mitochondria scattered throughout the cell section.  Functionally, mitochondria possess numerous enzymes necessary for aerobic metabolism in all cells.  This is especially true in muscle cells because of their high energy demands.  Smooth muscle, however, have considerably low phosphorylation capacity, therefore producing only one tenth the amount of adenosine triphosphate (per unit weight of muscle) relative to skeletal muscle (Stephens 2001). The Golgi apparatus in smooth muscle is involved in protein packaging and transport of proteins in addition to its role in several other synthetic and degradative processes in the cell.       9    Figure 2:  Organelles in a transverse section of an airway smooth muscle cell.  Major organelles of the airway smooth muscle include nucleus (N), mitochondria (M), rough endoplasmic reticulum (black arrowhead), and smooth endoplasmic reticulum (white arrowhead).  Rough endoplasmic reticulum can be identified by the large number of ribosomes in close proximity.  Other electron-dense structures within the airway smooth muscle include dense bodies (black circle) and dense plaques (white arrow).           10  1.2.3 Cytoskeleton  1.2.3.1 Organization of filaments Three different types of filaments have been identified in smooth muscle cells: thin, thick, and intermediate filaments.   A thin filament is an aggregate of proteins consisted of an actin filament backbone along with tropomyosin, caldesmon, and calponin.  Actin filaments are formed by the polymerization of G-actin monomers (a globular, 42 kDa protein) into two parallel strands rotated around the axis to form a double helix approximately 6-8 nm in diameter (Fig. 3) (Small and Sobieszek 1983).  Actin is present with four highly conserved isoforms in smooth muscle: two smooth muscle specific isoforms (α- and γ-actins), and two ubiquitously expressed non-muscle isoforms (β- and γ-cytoplasmic actins) (Vandekerckhove and Weber 1978).  Whether a thin filament is involved in contraction or cytoskeletal structure is believed to be determined by the actin isoform composition of the filament.  A unique characteristic that differentiates smooth muscle from skeletal muscle is the distribution of thin and thick filaments.  In smooth muscle, thin filaments are approximately ten times more prevalent, with a thin to thick filament ratio of ~20:1, as compared to 2:1 in skeletal muscle (Stephens 2001; Herrera, Martinez, and Seow 2004).  This abundance of thin filaments in smooth muscle is illustrated in Fig. 3 (black arrow). Thick filaments (or myosin filaments) are formed by the polymerization of class II myosin molecules with filament diameters ranging from 11-21 nm (Fig. 3).  Class II myosin molecules are found in non-muscle cells and smooth muscle cells in hollow 11  organs such as stomach, bladder, blood vessels and airways (Eddinger & Meer, 2007), and are able to self-assemble in solution to form side-polar filaments (Craig & Megerman, 1977; Xu et al, 1996).  The side-polar structure has also been seen in native myosin filaments isolated from skinned smooth muscle cells (Cooke et al, 1989) as well as in ultra-rapidly frozen, freeze-fractured and deep-etched intact smooth muscle cells (Hodgkinson et al, 1995).  As in striated muscle, cyclic interaction between myosin cross-bridges and actin filaments is the mechanism in smooth muscle contraction.  It is well known that the bi-polar myosin filaments in a striated muscle cell have the same length; whether this is true for smooth muscle is not clear.  The tail end of a smooth muscle myosin molecule has been shown to be important in filament self-assembly (Trybus & Henry, 1989).  It has been proposed that a tail-end to tail-end association of two myosin monomers is responsible for forming an anti-parallel dimer that serves as a nucleation site for filament growth (Cross et al, 1991).  An illustration of the type of filament assembly is shown in Fig. 4.  This type of filament growth may be governed by competing processes of polymer formation and dissolution that may not produce filaments of identical length, but may more likely produce filaments of variable lengths with a random distribution. Intermediate filaments (10 nm in diameter) in most smooth muscles are mainly composed of desmin (55 kDa) along with minor components of vimentin (Bagby et al. 1990).  They are one of the major elements of the cytoskeleton, encompassing approximately 15-25% of the total structural protein content in smooth muscle cells (Small and Sobieszek 1983).  There is evidence indicating intermediate filament association with dense bodies and dense plaques, making it a major contributor to the strength of the cellular cytoskeleton 12  (Zhang et al. 2010).  This is shown in Fig. 3 with numerous intermediate filaments (double arrows) located in close proximity to the electron-dense areas.  Intermediate filaments have also been shown to play essential roles in various cellular functions including signal transduction, force generation, and structural integrity (Helfand et al. 2005; Tang 2008).      13    Figure 3: Electron micrograph of transverse section of a tracheal smooth muscle cells.  Thin black arrows point to actin filaments; the thick white arrow points to a myosin filament; double black arrows point to intermediate filaments; the arrowhead points to a dense body; M, mitochondria.  (Modified from Liu et al. 2013)     14           Figure 4: Schematic illustration of a myosin monomer and a filament.  Represented by 8 myosin monomers arranged in an anti-parallel fashion characteristic of a side-polar filament.  (Liu et al. 2013)                14.5 nm 43.5 nm Monomer Filament 15  1.2.3.2 Dense bodies and dense plaques Dense bodies and dense plaques can be identified in electron micrographs of smooth muscle cells as distinct electron-dense foci throughout the cell.  The dense areas associated with the cytoplasm and membrane are called as dense bodies and dense plaques, respectively, and are both believed to serve as the smooth muscle equivalent of Z-disks in striated muscle.  More specifically, these dense structures are thought to act as anchoring sites for actin and intermediate filaments.  Functionally, this allows for contribution to both the transmission of force throughout the cell and also mechanical coupling with adjacent cells (Bagby et al. 1990).   Cytoplasmic dense bodies are mainly composed of α-actinin, but have also been shown to have minor components of calponin and β-actin (Geiger et al. 1981; North et al. 1994).  They are found as elongated structures scattered throughout the cytoplasm (Figs. 1 and 3) parallel to the longitudinal axis of the cell (Zhang et al, 2010).  Furthermore, this longitudinal orientation also positions the dense bodies so that they run parallel to the myosin filaments and the direction of force transmission (Herrera, Martinez, and Seow 2004).  Although very little is known about the structure of dense bodies in airway smooth muscle cells, studies have identified varying dense body dimensions in most visceral smooth muscle.  In these studies, dense bodies with lengths of up to 1.2 μm and widths of up to 0.3 μm were found (Gabella 1997).  In terms of function, Zhang et al. found that dense bodies could play a role in stiffness and plasticity in smooth muscle cells.  These intracellular cable-like structures are able to bear passive tension such that stretching the muscle cells in the relaxed state causes the cables to straighten.  Furthermore, shortening a muscle cell does not cause the cables to collapse.  The cables 16  are instead able to maintain tensile strength by quickly adapt to the new length and remain relatively straight.  These plastic structures may contribute to the ability of certain hollow organs to maintain stiffness over large length ranges (Zhang et al. 2010).   Dense plaques have much of the same protein composition as dense bodies but with the addition of vinculin and talin, due to its association with adherens junctions (McGuffee, Little, and Bagby 1989).  They are found as electron-dense bands along the cell membrane with an average width of 0.2-0.4 μm, 30-40 nm in thickness, and elongated over 1-2 μm in length (Fig. 1).  In smooth muscle, these bands have been suggested to provide mechanical coupling between myofilaments and the extracellular matrix by providing anchoring points for both actin and intermediate filaments (Stephens 2001).    1.2.3.3 Contractile apparatus – smooth muscle “sarcomere” Striated muscles possess a highly organized internal structure of repeating arrays of contractile filaments that appear as alternating light and dark bands (Fig. 5, left).  The basic contractile units in striated muscle are sarcomeres.  They are composed of overlapping actin and myosin filaments arranged in parallel and anchored to both the M-line and Z-discs by titin, which provides structural stability and also contributes to force transmission through the sarcomeres.   During a contraction, cyclic interactions between myosin cross-bridges and actin filaments allow for muscle shortening and force generation in a process described as the sliding filament mechanism (Huxley 1957). In smooth muscle, contraction is believed to be accomplished by the same sliding filament mechanism (Guilford and Warshaw 1998).  However, the internal contractile 17  elements are far less organized in smooth muscle cells and can be characterized by the lack of cross striations (Fig. 5, right).  Further advancement in the smooth muscle research has very much been hindered by the lack of knowledge pertaining to the structural organization of the contractile unit at the molecular level.  Recent studies, however, have found functional and structural evidence to support a model of the smooth muscle contractile unit.  Fig. 6 shows the proposed structure consisting of a side-polar myosin filament sandwiched by actin filaments on either side which are anchored to dense bodies.  During a contraction, cross-bridges (myosin heads) of the side-polar myosin filaments bind to the actin filaments on either side and pull them in opposite directions (Herrera et al. 2005; Syyong et al. 2011).      18   Figure 5: Structural comparison of skeletal and smooth muscle cells.  Left: Electron micrograph of a longitudinal section of mouse diaphragm (skeletal) muscle.  Contractile units are the highly organized structures known as sarcomeres, giving the muscle a “striated” appearance.  The organized myosin filaments are visible as a black band between the white lines.   Right: Electron micrograph of a longitudinal section of a sheep tracheal smooth muscle cell.  It is characterized by the lack of organized striation, but instead possesses more randomly distributed myosin filaments (white arrows).  (Liu et al. 2013)              19    Figure 6:  A proposed configuration of smooth muscle contractile unit.  Double arrows indicate the sliding direction of actin filaments.                               20  1.3  Smooth muscle contraction  Smooth muscle is distributed in the walls of various hollow organs, contributing to a wide range of functions of the organs such as the airways, stomach, blood vessels, and bladder.  They can be categorized as phasic (transient) or tonic (sustained) muscles depending on their speed and pattern of activation (Somlyo and Somlyo 1994).  Phasic muscles normally produce short-lasting but rapid contractions and are found in an array of organs including the uterus, urinary bladder, intestines, and stomach.  They are typically activated by neural stimulation or spontaneous action potentials that propagate from cell to cell.  In contrast, tonic muscles are rarely completely relaxed but instead produce gradual long-lasting contractions with relatively lower rates of energy consumption.  They are distributed throughout the walls of the airways and blood vessels and are typically activated by humoral or pharmacological agonists (Meiss 1997).           Experimentally, smooth muscle contraction can be induced isometrically or isotonically, depending on the variable that is kept constant.  Isometric contraction occurs when the muscle generates force while maintaining a constant length, whereas isotonic contraction occurs when length is allowed to vary while the muscle shortens against a constant load.  Muscle contractions in vivo, however, are rarely entirely isometric or isotonic.  Instead, they generally contract auxotonically with muscle shortening against a varying load. The mechanism for smooth muscle contraction involves a combination of chemical and physical interactions between receptors and contractile proteins, as illustrated in Fig. 7.  Muscle activation causes a rapid increase in intracellular [Ca2+]. This is caused by calcium influx from extracellular stores through the voltage-dependent and/or receptor-21  linked calcium channels (Frearson et al. 1976; Sobieszek 1977).  Calcium is also released from the sarcoplasmic reticulum via inositol 1,4,5-triphosphate (IP3) gated calcium channels through ryanodine receptors.  The cytosolic calcium is then able to bind to calmodulin, which in turn forms a complex with myosin light chain kinase (MLCK).  The activated kinase is then capable of phosphorylating myosin regulatory light chain (MLC20) leading to cross-bridge cycling and, therefore, muscle contraction.  However, a parallel antagonistic pathway involving myosin light chain phosphatase (MLCP) functions in dephosphorylating MLC20, thus counteracting the actions of MLCK.  Both the rho-kinase and protein kinase C (PKC) pathways are activated by G protein-coupled receptors and are known to have inhibitory effects on MLCP in smooth muscle (Somlyo and Somlyo 2004; Puetz et al. 2009).  More specifically, activated rho-kinase phosphorylates the myosin phosphatase-targeting subunit 1 (MYPT1), which directly inhibits the activity of MLCP.  Furthermore, both rho-kinase and PKC are able to phosphorylate phosphoprotein CPI-17, which also binds to and inhibits MLCP activity (Koyama et al. 2000; Dixon and Santana 2013).  Therefore the degree of muscle contraction that occurs when activated is dependent on the level of MLC20 phosphorylation, regulated by the relative activities of MLCK and MLCP.     Subsequently, muscle relaxation occurs when the intracellular [Ca2+] is decreased.  The removal of calcium can either be achieved by calcium extrusion out of the cell or by calcium uptake back into the sarcoplasmic reticulum.  The result is inactivation of MLCK, dephosphorylation of myosin regulatory light chain, and consequently termination of cross-bridge cycling (Pfitzer 2001).   22   Figure 7: Smooth muscle contraction pathway.  Ca2+ and  G protein-coupled receptor pathways in contraction and  relaxation.  Ach, acetylcholine; CaM, calmodulin; CPI-17, PKC-potentiated inhibitor protein (17kD); Gα, G alpha subunit; GPCR, G protein-coupled receptor; IP3, inositol 1,4,5-triphosphate; KCl, potassium chloride; LVGC, L-type voltage-gated Ca2+ channel; MLC20, myosin light chain (20kD); MLCK, myosin light chain kinase; MLCP, myosin light chain phosphatase; PKC, Protein kinase C; PLC, phospholipase C; ROCK, Rho kinase; RyR, ryanodine receptor; SR, sarcoplasmic reticulum.   23  1.4 Plasticity and length adaptation Hollow organs such as the bladder, stomach, and uterus undergo large changes in volume during normal physiological functioning.  The walls of these hollow organs are lined with smooth muscle, suggesting that the smooth muscle cells themselves are able to remain functional over substantial changes in length.  In smooth muscle, the ability of muscle cells to structurally and functionally adapt to the external stimuli can be categorized into two types, mechanical plasticity and phenotypic plasticity.  Mechanical plasticity refers to the muscle’s ability to quickly and efficiently alter the structure of its cytoskeleton and contractile apparatus in response to physical variations in the environment (C. Y. Seow and Fredberg 2001).  In contrast, phenotypic plasticity refers to a more drastic and long term process of adaptation.   This type of adaptation involves alterations in protein compositions and can be triggered by a wide array of stimuli that disrupt the homeostasis of the muscle (Halayko and Solway 2001).  In smooth muscle, the immediate effect of a change in muscle length (caused by externally applied force) is reduced force generation.  However, if the muscle is allowed to equilibrate at the new length, force generation can be recovered in a process called length adaptation (Bai et al. 2004).   Evidence for length adaptation has been reported in various types of smooth muscle cells.  Rabbit urinary bladder smooth muscles have demonstrated the ability to maintain their contractile ability even when lengthened to more than 7 fold their resting length (Uvelius 1976).  Similarly, studies on the malleability of the contractile apparatus in airway smooth muscle provide further evidence for mechanical plasticity (Chun Y. Seow, Pratusevich, and Ford 2000; Gunst et al. 1995).  Studies on swine carotid artery (Wingard, Browne, and Murphy 1995) and rat 24  anococcygeus muscle (Gillis, Cao, and Godfraind-De Becker 1988) have also shown similar levels of functional and structural plasticity.     25  CHAPTER 2:  Hypothesis, specific aims, and rationale   2.1  Hypothesis The general hypothesis guiding the thesis research is that within a smooth muscle contractile unit (sarcomere equivalent in smooth muscle) the myosin filaments do not have a constant length.  More specifically, within the actin filament lattice of the smooth muscle contractile unit, instead of a single myosin filament sandwiched by actin filaments, there are multiple myosin filaments of varying lengths lying tip-to-tip in series (Fig. 8).              26     Figure 8:  Hypothesized configuration of smooth muscle contractile unit with myosin filaments of variable lengths.  Double arrows indicate the sliding direction of actin filaments. 27  2.2  Specific aims The goal is to determine the length distribution of myosin filaments in serial transverse sections of smooth muscle cells.  To test our hypothesis, we performed the following experiments: 1) Smooth muscle tissue was fixed for electron-microscopy and serial ultra-thin (50-60 nm) cross-sections were obtained for electron microscopy.  Myosin filaments were traced through consecutive sections to determine the length of each myosin filaments.  Myosin filament lengths from various smooth muscle cell types were measured:  1) sheep airway smooth muscle; 2) sheep pulmonary arterial smooth muscle; 3) rabbit carotid arterial smooth muscle.  Rationale:  As mentioned in Chapter 1, formation of the side-polar myosin filaments in smooth muscle is a spontaneous process and there is no known mechanism that ensures that every filament formed is of a constant length.  Rather, polymerization and depolymerization of myosin filaments is likely determined by the affinity of the myosin dimers for each other as well as the propensity for polymerized dimers to dissociate from each other. If this is the case then a wide range of lengths for myosin filaments will be found.    2) The same protocol for fixation and electron microscopy was followed for intact airway smooth muscle activated with acetylcholine and serial ultra-thin (60 nm) cross-sections were obtained.  Activation of the muscle was induced by acetylcholine (10-4 M) and fixed at the plateau of isometric contraction.  Myosin filaments were 28  traced with the same method as in the relaxed smooth muscle to determine the length distribution within the sample.  Rationale:  It has been shown that phosphorylation of the myosin light chain facilitates myosin filament formation in solution (Trybus and Lowey, 1987).  There is also evidence that activation of smooth muscle leads to an increase in the mass of the myosin filaments (Kuo et al, 2003).  However it is not known whether the increase in the filament mass is due to an increase in the number of filaments or an increase in the average length of the filaments. Comparing measurements from relaxed and activated smooth muscle will allow us to assess the effect of light chain phosphorylated on filament formation.          29  CHAPTER 3:  Materials and methods  3.1  Tissue sample  Sheep lungs were obtained from a local abattoir; carotid arteries were obtained from euthanized New Zealand White rabbits; mouse diaphragms were obtained from euthanized C57 mice.  All protocols were approved by the Animal Care Committee of the University of British Columbia and conformed to the guidelines set out by the Canadian Council on Animal Care.     3.2  Tissue preparation  Tissues were place in physiological Kreb’s solution at 4°C immediately after removal from the animal.  Airway smooth muscle was obtained from sheep tracheal rings, with close attention paid to the in situ length of the trachealis prior to cutting of the C-shaped cartilage.  Tracheal rings were cut open on the ventral side through the C-shaped cartilage region.  A rectangular sheet of smooth muscle tissue, free of connective tissue and epithelial layer, was dissected from the tracheal ring.  The piece of tissue was then cut into multiple parallel strips along the longitudinal axis of the cell bundles.  Aluminum foil clips were then attached to both ends of the muscle strip for attachment to the force/length transducer.  Due to the size of the arterial smooth muscle layers, whole ring segments were used instead of strips.  Loose connective tissue was removed from the ring 30  segments, along with removal of the endothelial layer by gently rubbing the lumen side of the arteries with a pair of forceps.   The rings were then placed directly on to the force/length transducer.  The airway smooth muscle strips and ring segments of pulmonary and carotid arteries were then equilibrated in muscle baths containing physiological saline solution (Krebs’ solution) at 37°C for 1 h during which the muscles were stimulated electrically by electrical field stimulation for 10 s once every 5 min to attain maximal and stable isometric force production.  Electrical field stimulation was provided by a 60 Hz alternating current stimulator with platinum electrodes inside the muscle bath.  The Krebs’ solution with pH 7.4 at 37°C was aerated with a gas mixture (carbogen) containing 5% CO2 - 95% O2, and a composition of 118 mM NaCl, 4.5 mM KCl, 1.2 mM NaH2PO4, 22.5 mM NaHCO3, 2 mM MgSO4, 2 mM CaCl2 and 2 g/l dextrose.  The muscle was considered equilibrated when it developed stable maximal isometric force with negligible resting tension. The tissues were then fixed at in situ length for electron microscopy.  For tissues fixed in the activated state, acetylcholine (10-4 M) was used for stimulation and the muscle was fixed at the plateau of isometric contraction about two min after onset of stimulation.  Only tracheal smooth muscle strips were fixed in the activated state in this study.  More details regarding tissue dissection and preconditioning can be found in previous publications from our laboratory  3.3  Electron microscopy  Primary fixation for 15 min was done while the muscle preparations were still attached to the apparatus that measured muscle force.  The fixing solution contained: 1% 31  paraformaldehyde, 2.5% glutaraldehyde, and 2% tannic acid in 0.1 M sodium cacodylate buffer.  The fixing solution was pre-warmed to 37°C.  After primary fixation, the muscle strip was removed from the apparatus and cut into small blocks (~2 × ~0.5 × ~0.2 mm in dimension) in cold fixation buffer and kept in the same fixative for 2 hours in an ice bucket on a shaker.  During the secondary fixation the tissue blocks were first transferred to 1% osmium buffer for 1.5 hours at ice temperature on the shaker, followed by three washes with 1.25% NaHCO3 aqueous solution and then distilled water (10 minutes per wash).  The tissue was stained with 1% uranyl acetate, dehydrated with increasing concentrations of ethanol (50%, 70%, 80%, 90%, 95%, 100%) and then propylene oxide.  After dehydration, small tissue pieces were embedded in resin (TAAB 812 mix).  The resin blocks were sectioned with a diamond knife.  Transverse sections were cut at 50 nm thickness for tracheal smooth muscle fixed in the relaxed state, and sections were cut at 60 nm thickness for other smooth muscle preparations.  For diaphragm (skeletal) muscle, sections were cut at 100 nm thickness.  The sections were picked up on Formvar covered slotted grids.  The sections were further stained with 1% uranyl acetate and lead citrate.  Images of transverse and longitudinal sections of muscle cells were obtained with an electron microscope (Philips/FEI Tecnai 12 TEM) equipped with a digital camera (Gatan 792) at a magnification of 37,000X.  To capture the whole cross-section of a single cell it often required taking multiple images of a single cell and merging them together with the help of Adobe Photoshop.   32  3.4  Image analysis  Adobe Photoshop (CS6) and Fiji (a modified version of ImageJ 1.47q) were used to facilitate measurements of myosin filament length.  In serial sections myosin filaments were traced by superimposing consecutive sections utilizing the layer visibility function (Adobe Photoshop).  The length of a filament measured in a series of transverse sections is calculated as the number of sections in which the same filament was present times the section thickness (e.g., 50 nm for tracheal smooth muscle sections).  In longitudinal sections the myosin filaments were measured using a line function in Fiji.   3.5  Myosin filament length measurement  Figs. 1, 2 and 3 all show different cross-sections of sheep tracheal smooth muscle cell.  Three series of consecutive cross-sections from 3 cells were obtained from 2 sheep tracheas.  Each series contained at least 69 sections. The section thickness for tracheal smooth muscle was 50 nm.  Two series of consecutive cross-sections from 2 pulmonary arterial smooth muscle cells were obtained from a sheep.  Each series contained at least 30 sections.  Two series of consecutive cross-sections from 2 carotid arterial smooth muscle cells were obtained from a rabbit.  Each series contained at least 53 sections.  The section thickness for arterial smooth muscles was 60 nm.  Fig. 9 shows an example of serial sections (sheep tracheal smooth muscle) used in the filament length measurements.  "Landmarks" such as mitochondria, dense plaques, dense bodies, microtubules and thick filaments were used to register serial sections and thus help in tracing a particular thick 33  filament from section to section.  The "landmarks" moved slightly from one section to the next, and they appeared and disappeared quite frequently throughout the series.  We therefore could not use the same "landmarks" throughout a long series of sections; instead, we relied more on the same structures identified in the sections immediately above and below as a guide to continuously modify the section registration as we worked through the series.  In the example shown in Fig. 9, the consecutive sections (each showing a small area within the serial cell cross-sections) are labeled 1-11, and each section is subdivided into 25 squares by a grid labeled A to Y.  Two microtubules, one near grid K and the other near grid P, (Fig. 9, lower middle panel, double arrows) were used as one of the "landmarks" that enabled us to follow the myosin filaments from one section to the next throughout the whole series.  The other guidance used was the positions of the surrounding myosin filaments with respect to the filament we were tracing.  The (surrounding) myosin filaments in grid G, L and R can be followed from section 1 to 4; the filament in M (the one being traced) appears in section 2, and disappears in section 7.  The length of that filament was calculated as the number of contiguous sections containing the filament multiplied by the section thickness, that is, 5 x 50 = 250 nm in this example.  Only the filaments whose appearance and disappearance in the serial sections could be clearly identified were included in the length measurement.  Because of the discrete nature of the measurements using serial cross-sections with finite thickness the filament length determined by this method actually represented a range of lengths.  For example, the number of filaments in the 50-nm bin would contain any observable filaments up to 50 nm in length and in the 100-nm bin the filament lengths would range from 50-100 nm, and so forth.   34  Figure 9: Measurement of myosin filament length in serial sections of electron micrographs.  Eleven consecutive sections are used in this demonstration, labeled 1-11 as shown in the bottom left panel.  For each section, the area is subdivided into 25 squares (yellow grid) and labeled A to Y as shown in the bottom right panel. In the bottom middle panel, thin arrows point to actin filaments, thick arrows point to myosin  filaments (surrounded by actin filaments), and double arrows point to microtubules. Details for following a thick filament through the serial sections are described in the text (Liu et al. 2013).   35  3.6  Statistical analysis  Two-way analysis of variance (ANOVA) was used for comparison of filament length distribution between groups. This examines the effect of two categorical independent variables on a single continuous dependent variable.  N was the number of cells used in cross-sectional tracing of filaments.  Data are presented as means± standard error of means (SE) unless otherwise specified.  Statistical significance corresponds to P<0.05.  All statistical analyses were carried out in SigmaPlot 11. The Bland-Altman method was used to evaluate the inter-observer differences in the measurement of filament length using serial cross-sections.  This is a statistical method for assessing the agreement between two methods of measurement.  The resulting bias corresponds to the degree of systematic variation between measurements.    36  CHAPTER 4: Results   4.1  Control observations  Mouse diaphragm (skeletal) muscle was used to test whether we could accurately determine the skeletal myosin filament length in both longitudinal sections and serial cross-sections.  Fig. 5 shows a longitudinal section of the skeletal muscle.  The length of the myosin filaments measured from the longitudinal sections was ~1.15 µm.  Fig. 10 shows 8 cross-sections (a-h) taken from a series of 13 consecutive sections of 100 nm thickness.  Note that not all consecutive sections are shown in Fig. 10.  The serial section numbers corresponding to the panels (a-h) in Fig. 10 are: a(1), b(2), c(4), d(6), e(8), f(11), g(12), h(13).  Myosin filaments are present from section 2 to 12 inclusive, indicating that the filament length is about 1.1 µm, in good agreement with the measurements from longitudinal sections.   37    Figure 10: Serial cross sections of mouse diaphragm (skeletal muscle).  Selective sections from a 13-section series of consecutive cross-sections from the same mouse diaphragm shown in Fig. 5. Each image size is 1.1 x 1.1 µm.  (Liu et al. 2013)   38  4.2  Inter-observer differences   At the beginning of this study we assessed the inter-observer differences in the measurement of filament length using serial cross-sections to determine how much influence the subjective recognition of filament pattern had on the results.  Fig. 11A shows the filament length distribution measured by two observers from a 26-section series from a sheep trachealis.  The Bland-Altman plot (Fig. 11B) shows no systematic variation from one observer to the other.  The length measurements included in the rest of this study were all carried out by one observer (Observer 1, JL).     39  Filament Length (um)0.0 0.2 0.4 0.6 0.8 1.0Observation Frequency0.000.050.100.150.200.250.300.35Observer 1Observer 2                                      Figure 11: Inter-observer difference in the measurement of myosin filament lengths.  A) The distribution of observation frequency with filament length for Observer 1 (JL) and Observer 2 (CS).  The total number of filaments traced by observers 1 and 2 are 593 and 332 respectively.  B) A Bland-Altman plot for the data in panel A. The small bias indicates no systematic variation with the measurements from the two observers.  (Liu et al. 2013)  Observation Frequency0.00 0.05 0.10 0.15 0.20 0.25 0.30% Difference/mean (Observer 1 - Observer 2)-100-80-60-40-20020406080100Bias = -1.07 %40  4.3  Mathematical model for linear polymerization and fragmentation  To help interpret observations from experiments regarding myosin filament polymerization a mathematical model was developed to describe the polymerization process.  Because myosin filaments are un-branched, we assume that they are formed within actin filament lattices by a dynamic process of linear aggregation and fragmentation.  The basic units for myosin polymerization could be myosin dimers.  However, we define the basic polymerization unit as the shortest quantifiable myosin filament in serial sections cut for transmission electron microscopy (EM).  For section thickness of 50 nm, a filament segment perpendicular to the cross-section and exposed on both surfaces of the section is probably an oligomer containing at least 4 dimers in series (Fig. 4).  For section thickness of 60 nm, the filament probably contains 5 dimers in series.    For the mathematical description we consider a simple linear one dimensional lattice polymerization model where at each “time step” bonds are formed between the basic polymerization units.  We assume that polymerization of linear polymers involves two competing processes: aggregation (formation of linear bonds between neighboring polymerization units) and fragmentation (breaking of the bonds), which we will designate as p and q, respectively.  The two processes occur independently and randomly but with certain probabilities.  Filament formation in this model is therefore a dynamic process which when given enough time will settle into a steady state.  Filament length distribution in a muscle cell fixed in the relaxed state or at the plateau of an isometric 41  contraction represents such a steady state distribution.  For the mathematical description we consider a simple linear one dimensional lattice polymerization model where at each “time step” bonds are formed between the basic polymerization units with probability p and are broken with probability q.  For a total of Nmax sites an equation describing the evolution of the mean number of n bonds reads   nqpnNdtdn  max The first term describes the formation of bonds at (Nmax − n) possible sites and the second term is just the decay of existing bonds.  For q > 0 this equation has a dynamic equilibrium or steady state where the right hand side vanishes, dn/dt = 0, and the population is constant, n ≡ neq.  From this condition we obtain the mean probability r for a bond to be intact, qppNnr eq  max    [1a] Next we examine the distribution of cluster sizes (or filament lengths) in this dynamic equilibrium.  Since all probabilities are assumed to be independent, we are simply considering a line of sites or bonds that are occupied with probability r.  Note that r depends on a combination of p and q but is really the only relevant parameter here.  The probability to observe a cluster (filament) of l connected sites is proportional to rl, lrlP )(  This behavior is also found in a class of problems called “percolation” (Bunde & Havlin, 1996).  Since rl = el·ln(r), this equation is equivalent to a pure exponential decay 42  /)( lelP       [2a] with parameter λ = −1/ln(r).  λ is therefore the length constant in the exponential decay length distribution.  The mean probability r can be expressed as a function of λ. 1 er       [3a] Also because dle l  0 /  the normalized length distribution becomes: /)(lelP     [4a]  Note that the validity of the model (Equation 4a) is subject to the conditions described above.    4.4  Myosin filaments in relaxed airway smooth muscle  Fig. 12 shows the observation frequency as a function of myosin filament length in the relaxed sheep tracheal smooth muscle.  There were more short filaments observed, and the observation frequency decreases as the filaments get longer.  The dashed line in Fig. 12 shows the exponential fit to the data.  The exponential equation used is of the form: OF = Ae-L/Lo, where OF is the observation frequency, A is the initial value when L = 0, L is the filament length in μm, and Lo is the length constant.  The following values were 43  obtained from the exponential fit shown in Fig. 12: A = 0.2799±0.0096, 1/Lo = 5.498±0.2299 (μm-1), the goodness of fit is 0.9754.  The length constant Lo for the distribution shown in Fig. 12 is therefore ~0.182 μm.  This is the length where the observation frequency is about 37% of the initial value, and is used to indicate how fast the frequency decays.  Dividing Lo by the length of the basic unit for polymerization, 50 nm in this case, we obtained the dimensionless length constant λ (λ = 0.182/0.05 = 3.64).  λ is related to p and q (see Eqn. 1), but without further information on the dynamic process of filament polymerization it is not possible to determine the values of p and q separately.  The mean probability for the basic polymerization units to link up with one another can be calculated from Equation 3 to give r = 0.760.    44  Filament Length (m)0.0 0.5 1.0 1.5 2.0 2.5Observation Frequency0.000.050.100.150.200.25  Figure 12: Distribution of observation frequency over myosin filament lengths measured through tracing of consecutive serial cross-sections (50 nm thickness) of sheep tracheal smooth muscle cells (circles). A total of 2627 filaments in 3 cells from 2 tracheas were traced in this group of measurement.  Means and standard errors are plotted.  Dash-line shows an exponential fit (See text for details of simulation model).   (Liu et al. 2013)   45  4.5  Myosin filaments in relaxed arterial smooth muscle  Fig. 13 shows frequency distribution of myosin filament length in smooth muscles from sheep pulmonary artery and rabbit carotid artery in the relaxed state.  The length distribution data from the two types of muscles are very similar and therefore they were plotted together as a group.  The dashed line shows an exponential fit (OF = Ae-L/Lo) with A = 0.6808±0.0051, 1/Lo = 8.623±0.0693 (μm-1), and goodness of fit 0.9998.  The length constant (Lo) for the distribution shown in Fig. 13 is therefore ~0.116 μm.  To convert Lo to λ we divided Lo by the length of the basic polymerization unit, 60 nm in this case, to obtain λ = 1.93, indicating a faster decay of the length distribution when compared to that of relaxed tracheal smooth muscle (λ = 3.64, Fig. 13).  The mean probability for polymerization (r) was calculated from Equation 3 as 0.596.  Two-way ANOVA indicates that the distributions in relaxed tracheal smooth muscle and relaxed arterial smooth muscles (shown in Fig. 12 and Fig. 13 respectively) are significantly different (p<0.01) 46  Filament Length (m)0.0 0.2 0.4 0.6 0.8 1.0 1.2Observation Frequency0.00.10.20.30.40.5  Figure 13: Distribution of observation frequency over myosin filament lengths measured through tracing of consecutive serial cross-sections (60 nm thickness) of arterial smooth muscle cells.  Four groups are plotted representing 4 cells, 2 from sheep pulmonary arterial smooth muscle (open symbols) and 2 from rabbit carotid arterial smooth muscle (solid symbols). The number of filaments traced for the groups are: 2335 (   ), 1639 (   ), 1379 (   ), and 756 (   ).  Dash-line shows an exponential fit (See text for details of model simulation).    (Liu et al. 2013)   47  4.6  Filament length measured in longitudinal sections  Fig. 5 shows an example of a longitudinal section of sheep airway smooth muscle in the relaxed state. Myosin thick filaments could be identified and their length measured.  Fig. 14 shows the results of measurements from longitudinal sections (circles).  These results were compared with those measured in serial sections (solid line, re-plotted using mean values from Fig. 12).  Note that for filament lengths greater than 0.15 µm the measurements made on longitudinal sections consistently underestimate the filament length when compared with the corresponding lengths measured from serial sections.  This is likely because the myosin filaments are not perfectly in parallel with the surface of the longitudinal section (which is only 50 nm thick).  The likelihood for the filament exiting the plane of the longitudinal section increases with increasing length of the filament.        48   Figure 14: Comparison of myosin filament length distributions measured from tracing serial cross-sections (solid line) and from longitudinal section with direct length measurement (circles) in tracheal smooth muscle cells.  Data for the solid line plot are mean values from Fig. 12.  For measurements in longitudinal section, the lengths of 3848 filaments were measured in 3 different cells.   (Liu et al. 2013)  Filament Length (m)0.0 0.5 1.0 1.5 2.0 2.5Observation Frequency0.000.050.100.150.200.2549  4.7  Myosin filaments in activated smooth muscle   Measurements of myosin filament length in the activated state were carried out in only one type of smooth muscle – sheep tracheal smooth muscle.  The muscle strips were fixed at the plateau of isometric contraction induced by acetylcholine (M-4) at 37°C.  Fig. 15 shows the observation frequency (solid circles) as a function of myosin filament length.  Means and standard errors are plotted.  The solid line shows an exponential fit (OF = Ae-L/Lo) with A = 0.6064±0.0123, 1/Lo = 7.861±0.1756 (μm-1), and goodness of fit 0.9977.  The length constant (Lo) is therefore ~0.127 μm.  To convert Lo to λ we divided Lo by the length of the basic polymerization unit, 60 nm in this case, to obtain λ = 2.12  indicating a faster decay in the distribution for the activated tracheal smooth muscle compared with that of the relaxed one (λ = 3.64, Fig. 12).  The mean probability for polymerization (r) was calculated from Equation 3 as 0.624, which is also lower compared with the r value (0.760) for the relaxed tracheal smooth muscle.  Two-way ANOVA indicates that the filament distribution patterns in the relaxed (Fig. 12) and activated (Fig. 15) tracheal smooth muscles are significantly different (p<0.0001).  Bonferroni multiple comparisons shows significant difference (p<0.001) between the two groups of data at lengths ≤ 0.2 μm, but no significant difference at other lengths. 50   Filament Length (m)0.0 0.2 0.4 0.6 0.8 1.0 1.2Observation Frequency0.00.10.20.30.4  Figure 15:  Distribution of observation frequency versus myosin filament lengths measured in ACh-activated sheep tracheal smooth muscle cells (solid circles). A total of 7851 filaments in 4 cells from 2 tracheas were traced in this group of measurement.  Means and standard errors are plotted.  Solid line shows an exponential fit. (See text for fitting parameters).  The dashed-line and open circles show the length distribution of myosin filaments in relaxed tracheal smooth muscle, re-plotted from Fig. 12 for comparison.  The two curves cross over at about 0.3 µm. Compared with relaxed muscle, the activated muscle has more short (<0.3 µm) filaments and less long (>0.3 µm) filaments.  (Liu et al. 2013)  51  CHAPTER 5:  Discussion   The most important findings from this thesis research is that myosin filaments in both relaxed and activated smooth muscle do not have a constant length, and that the median length of myosin filaments in activated smooth muscle is shorter than that in relaxed smooth muscle. The observed increase in myosin filament mass in activated smooth muscle  is therefore not due to an increase in average length of the myosin filaments but due to formation of shorter but more numerous filaments (Kuo et al. 2003; Godfraind-De Becker and Gillis 1988; Xu et al., 1997; Smolensky et al. 2005; Gillis et al., 1988).    5.1  Discrepancies with previous studies on myosin filament length  The structural organization of the contractile filaments in smooth muscle is not well understood (Seow, 2005a).  A smooth muscle contractile unit functionally analogous to a sarcomere in striated muscle has not been clearly delineated.  In striated muscle, determining the lengths of thin (actin) and thick (myosin) filaments was an important step in elucidating the precise structure of the sarcomere.  Myosin filaments in smooth muscle have been estimated to have lengths of 2-2.2 µm (Ashton et al, 1975; Herrera et al, 2005).  Examination of many published electron micrographs of longitudinal section of smooth muscle cells and hundreds of unpublished micrographs from our own laboratory revealed a wide range of myosin filament lengths.  This could be due to variations in the orientation of the filaments with respect to the surface of the section, because only when 52  a filament runs perfectly or nearly perfectly parallel with the section will its whole length be embedded within the relatively thin section.  However, it could also be due to myosin filaments of smooth muscle simply not having a constant length, contrary to their counterparts in striated muscle.  To resolve these possibilities we used ultra-thin (50-60 nm) serial transverse sections of smooth muscle cells to map out individual myosin filaments and trace the filaments from section to section to determine their lengths.  The control observation (Figs. 5 and 10) shows that for well-aligned filament arrays such as those found in mouse diaphragm muscle both longitudinal and serial cross-sectional measurements agree with each other.  However, for filaments not perfectly aligned (Figs. 5, 9 and 12), under-estimation of filament length is likely to occur in longitudinal measurements.  For this study we chose the method of tracing filaments in serial cross-sections.  The same approach was taken by Ashton et al (1975).  They concluded from their study that myosin filaments had a uniform length of about 2.2 µm.  This is fundamentally different from the conclusion of the present study.  The discrepancy could be due to the following reason.  Ashton and colleagues used relatively thick sections (400-500 nm) which are almost 10 times thicker than our serial sections.  This effectively reduced the resolution of their measurements.  We regularly detected 50-150 nm gaps in serially (end-to-end) aligned individual filaments, these gaps would likely be missed in 500 nm thick sections and therefore short but linearly aligned filaments could mistakenly be identified as a long filament.      53  5.2  Proposed model of a smooth muscle contractile unit  It is possible that the actin filament lattices have a uniform dimension but the myosin filaments within the lattices possess variable lengths, as illustrated in Fig. 16.  The side-polar nature of smooth muscle myosin filaments allows a myosin dimer to act as a ratchet when activated, pulling the actin filament in opposite directions.  Whether these dimers link themselves together to form a filament or not will not affect how they interact with actin filaments to produce force, if the contractile unit in smooth muscle is indeed constructed as that illustrated in Fig. 16.  This is analogous to a group of men standing in a straight line pulling two ropes in opposite directions with their hands.  Whether these men are chained together or not should not affect their performance.  If this is the case then there is no need for myosin filaments in smooth muscle to have a uniform length as required in striated muscle.  The existence of filamentous myosin in smooth muscle may be due to random interactions between myosin dimers in close proximity, as suggested by our mathematical simulation.  Another unique feature of smooth muscle is that within the cells there is a significant pool of monomeric myosin molecules that could be used as substrate for polymerization (Milton et al, 2011).  Such myosin reserve has never been identified in striated muscle.  In Fig. 16 the substrate (myosin dimers) for polymerization is assumed to exist both outside and within the actin filament lattice.  As outlined in the mathematical model presented in the Results section and in Chapter 5.2 with mode details, the probability of a dimer within the actin filament lattice to link up with a neighboring dimer is p, and a link between dimers has a probability q to break.  Under a dynamic equilibrium the mean probability for a link to exist between dimers is r which is 54  equivalent to p/(p+q) (Equation 1).  A snapshot of the dynamic equilibrium is provided by a fixed cell.  Unfortunately we cannot resolve the individual values for p and q, but only the mean probability r from curve fitting.  More information about how the dimers interact with each other within the actin filament lattice in vivo is needed to resolve the p and q values.        55   Figure 16: Schematic illustration of an actin filament lattice containing myosin filaments with variable lengths. Myosin monomers and dimers are assumed substrates for myosin filament formation and they are assumed to be present both inside and surrounding the filament lattice.  (Liu et al. 2013)  56  5.3  Facilitation of myosin filament polymerization in vivo  It is known that phosphorylation of the regulatory myosin light chain facilitates myosin filament formation in solution (Suzuki et al, 1978; Smith et al, 1983; Cross et al, 1986; Kendrick-Jones et al, 1987; Trybus & Lowey, 1987).  It is therefore possible that the myosin filament length distribution could be altered in activated smooth muscle.  Fig. 15 shows that the distribution was indeed altered: the distribution curve shifted significantly to the left indicating that the average length of activated myosin filaments was shorter compared with that of the relaxed ones.  This is reflected in the change in the mean probability for polymerization (Equation 3) from that in the relaxed state (r = 0.760) to that in the activated state (r = 0.624).   Ip et al (2007) observed in solution that although more myosin filaments were formed when the myosin molecules were phosphorylated, the average length of the activated filaments was shorter, in agreement with the present observation.  It should be pointed out that in solution the ionic strength has to be low (<100 mM) for the filaments to self-assemble, and also the length distribution resembles a bell-shape rather than an exponential decay pattern (Ip et al, 2007).  The in vitro mechanism of filament formation at low ionic strength is likely different from the in vivo mechanism.  At physiological intracellular ionic strength of ~200 mM and ~5 mM [MgATP], little if any myosin filaments could exist in solution (Craig et al, 1983).  The fact that we observed myosin filaments in intact muscles suggests that the intracellular environment somehow facilitated the polymerization.  Some possible mechanisms have been suggested. Mahajan et al (1989) have observed greatly accelerated (5-fold) myosin filament assembly in the presence of actin filaments, and a key step to myosin filament 57  formation appears to be the transient initial binding of myosin monomers to actin filaments.  Applegate and Pardee (1992) have also provided evidence for myosin filament formation facilitated by filamentous actin.  Katayama et al (1995) have demonstrated that at physiological levels of [MgATP] disassembly of unphosphorylated myosin filaments occurs; however, reassembly of the filaments occurs when caldesmon is added. Telokin, also known as kinase related protein (KPR), has also been shown to help maintain myosin filaments in relaxed smooth muscle cells (Kudryashov et al, 2002).  The finding that myosin filaments in the relaxed state on average are longer than those in the activated state (Fig. 12 and Fig. 15) may be explained by the filament stabilizing effect of some proteins like caldesmon (Sutherland & Walsh, 1989; Kudryashov et al, 2002) and 38k protein (Okagaki et al, 2000).     5.4  Implications of the mathematical  model for linear polymerization  The fact that a simple linear polymerization model could be used to simulate myosin filament polymerization to produce an exponential length distribution suggests a possible molecular mechanism governing the polymerization process.  The model suggests that the propensity for myosin dimers to form filaments is primarily governed by the binding affinity (related to p) between the dimers and the stability of the bonds (related to q) between them.  Unlike in striated muscle, the mechanism for ensuring that all myosin filaments have the same length is likely absent in smooth muscle.  In Appendix I an alternative model for filament polymerization is explored.  In this linear polymerization 58  model it is assumed that myosin filaments can only lengthen or shorten by adding or subtracting dimers at the two ends of a filament and that no fragmentation can occur by breaking bonds in the mid segment of the filament.  As shown in the model this type of filament growth leads to a normal distribution of the filament length centered on a single length determined by the probabilities associated with addition and subtraction of dimers to and from the ends of the filament.  This is very different from the exponential decay pattern and therefore cannot be used to explain the present data.    It is interesting that both pulmonary and carotid arterial smooth muscle possess very similar length distributions for their myosin filaments (Fig. 13).  This suggests that the myosin isoforms in mammalian arterial smooth muscles may form dimers that are very similar in binding affinity and bond stability.  The difference in the distribution between arterial and tracheal smooth muscle (Figs. 12 and 13) on the other hand suggests that the binding affinity and bond stability are different in myosins from smooth muscles with a greater difference in embryonic origins.  The mean probability for polymerization for tracheal smooth muscle (r = 0.760) is greater than that for arterial smooth muscle (r = 0.596).  In other words, compared with tracheal smooth muscle, arterial smooth muscle possesses myosin isoforms with less likelihood to form long filaments.        59  5.5  Limitations of the measurement technique   It should be pointed out that myosin filaments shorter than 50 nm (approximately the dimension of an octomer depicted in Fig. 4) may not be recognized as a filament in our measurements; and it is highly likely that filaments shorter than 50 nm do exist.  The “backbone” of a myosin filament is made up of the tails of myosin molecules. The length of a myosin molecule from head to tail is about 150 nm (Elliott et al, 1976), and about 100 nm of the tail is bundled into the backbone. The distance between myosin heads on a filament (Fig. 4) is about 14.5 nm and that means the backbone of a long filament contains multiple (more than two) tails bundled in parallel.  On the other hand, the backbone of a dimer contains only two tails.  The cross-sectional profile of a dimer backbone will therefore be thinner than that of a long filament and it will not likely be recognized as a thick filament in our method of measurement.  The short filaments (myosin hexamers, tetramers, and dimers) may therefore be “invisible” to our method of detection and unaccounted for.  There is no reason to suggest that these short filaments could not interact with actin filaments and participate in force generation.  Caution therefore must be taken in calculating force generated per myosin filament length (or cross-bridge) based on EM images, because of the possible existence of unaccounted short myosin filaments.        60  5.6  New role for myosin in smooth muscle length adaptation  A property that distinguishes smooth from striated muscle is the structural plasticity associated with the former (Pratusevich et al, 1995; Gunst et al, 1995; Bursac et al, 2005; Seow, 2005b).  The lack of stable myosin filaments with uniform length may allow smooth muscle to redistribute myosin molecules more easily in its adaptation to large changes in cell geometry that requires restructuring of the cell’s cytoskeleton and contractile apparatus.    5.7 Final conclusion  Myosin filaments in both relaxed and activated smooth muscle do not have a uniform length and analysis suggests that the distribution of filament length is a result of a dynamic equilibrium between polymerization and de-polymerization of myosin molecules driven by predictable probabilities of the myosin dimers to bind with and dissociate from each other.    61  Bibliography  Anderson, Richard G. W. 1998. “The Caveolae Membrane System.” Annual Review of Biochemistry 67 (1): 199–225. doi:10.1146/annurev.biochem.67.1.199. Applegate, D., and J. D. Pardee. 1992. “Actin-Facilitated Assembly of Smooth Muscle Myosin Induces Formation of Actomyosin Fibrils.” The Journal of Cell Biology 117 (6): 1223–30. Ashton, F. 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Murphy. 1995. “Dependence of Force on Length at Constant Cross-Bridge Phosphorylation in the Swine Carotid Media.” The Journal of Physiology 488 ( Pt 3) (November): 729–39. Xu, J. Q., J. M. Gillis, and R. Craig. 1997. “Polymerization of Myosin on Activation of Rat Anococcygeus Smooth Muscle.” Journal of Muscle Research and Cell Motility 18 (3): 381–93. Xu, J. Q., B. A. Harder, P. Uman, and R. Craig. 1996. “Myosin Filament Structure in Vertebrate Smooth Muscle.” The Journal of Cell Biology 134 (1): 53–66. Zhang, Jie, Ana M. Herrera, Peter D. Paré, and Chun Y. Seow. 2010. “Dense-Body Aggregates as Plastic Structures Supporting Tension in Smooth Muscle Cells.” American Journal of Physiology - Lung Cellular and Molecular Physiology 299 (5): L631–38. doi:10.1152/ajplung.00087.2010.   67  Appendix I  Mathematical description for linear polymerization by extension at the two ends   Here we assume polymerization of linear polymers only by adding or subtracting polymerization units at the two ends of a polymer with no fragmentation in the middle.  Assumptions associated with the model are: 1) Myosin polymerization starts with a basic polymerization unit – a dimer (or nucleation site) and the filament length (L) increases or decreases due to addition or subtraction of dimers only at the ends of a filament. 2) Once the bonds are made between two dimers they will not break (or fragment) except for those at the two ends of a filament.  3) The probability for adding dimers at either end of a filament is p and for removing them is q.  4) There is a large number of nucleation sites but the number of dimers surrounding any particular nucleation site is finite and that when all available dimers surrounding a nucleation site are polymerized the final myosin filament length associated with that particular site is Lmax.  This leads to the following equations describing the length (L) of a particular filament associated with a particular nucleation site: qpLLdtdL  )( max    [1b] Let l = L/Lmax, we have qpldtdl  )1(    [2b] 68  At equilibrium (dl/dt = 0),  pqleq 1     [3b] where leq is the length of a particular filament at a particular site at equilibrium.  The model (Equation 3b) therefore predicts a unique filament length when there is a unique q/p ratio.  Now let us examine the distribution of leq for a large number of nucleation sites.  If there is no variability in p, q, and the number of available dimers surrounding a site, then there will be a single length distribution, i.e., leq.  If there is variability, for example, in the number of available dimers associated with different nucleation sites that follows a normal distribution then the filament length distribution will have a normal distribution with a peak occurring at leq. Note that the peak distribution is distinctively different from the exponential distribution associated with the model described in Chapter 5.2.  The model described here therefore can be excluded as a candidate to explain the data presented in the main text. 

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