Thomas Dean Pollard, M.D. - US grants

The proposed experiments are designed to identify the energy transducing enzyme(s) for mitosis and other microtubule-dependent movements through microinjection into living cells of active site specific monoclonal antibodies to various myosins and dyneins, to characterize the molecular interactions responsible for actin filament binding to micro-tubules and to determine the morphological relationships among actin, myosin and micro-tubules in the mitotic spindle and actin and myosin in the contractile ring. We will complete the characterization of monoclonal antibodies to Acanthamoeba myosins-I and II, identify the mechanism by which some interfere with the interaction of purified actin and myosin, use the antibodies to establish that antagonists have been identified for all classes of myosin in the cell and then microinject individual and combinations of these active site specific antibodies into dividing cells to test which myosins (if any) are required for cytokinesis and mitosis. Identical studies will be carried out using human cytoplasmic myosins isolated from platelets to elicit and test monoclonal antibodies that will then be microinjected into dividing HeLa cells. We will complete the characterization of our collection of monoclonal antibodies to sea urchin cytoplasmic dynein in collaboration with Drs. Begg and Pratt (Harvard Medical School) and test them for effects on mitosis by microinjection. We will raise and characterize monoclonal antibodies to Tetrahymena ciliary dynein. They will be used for studies in collaboration with Dr. Ken Johnson (Penn State University) on the mechanism of dynein and to test by microinjection whether protiens with active sites similar to ciliary dynein are required for mitosis. The biochemical studies on actin-microtubule interaction will focus on how the microtubule associated proteins cross-link the two fibers and how this interaction is regulated by phosphorylation and other factors. We will study by electron microscopy the distribution of actin filaments and myosin in dividing HeLa cells using ferritin-labeled monoclonal antibodies to myosin and 2 improved fixation techniques for actin filaments.

Our long range goal is to understand the molecular mechanisms by which actin and its associated proteins participate in cellular movements and the structure of cytoplasm. We will use Acanthamoeba as our model system. To clarify the mechanism of actin polymerization, we will characterize nucleotide hydrolysis during polymerization, reconcile the mechanism of elongation that we deduced from measurements in the electron microscope with observations on bulk samples, analyze the mechanism of nucleation by direct methods, characterize the dynamics of actin filaments at seady state and study the mechanisms of action of cytochalasin and phalloidin. To understand the functions of the actin monomer binding protein called profilin, we will determine the 3-D structure of Acanthamoeba profilinI, profilin-II and ventebrate profilin at atomic resolution by X-ray crystallography, identify amino acid contacts between profilin and actin by chemical crosslinking and protein sequencing so that we can establish how the proteins bind together, characterize the dynamics of the binding of profilin to actin monomers and filaments and search for factors that might regulated this interaction, localize the 2 profilin isoforms in Acanthamoeba and characterize a library of monoclonal antibodies to profilins with the goal of identifying inhibitors of actinprofilin interaction for use in other experiments. Actophorin is an actin monomer binding and filament severing protein. We will determine the primary structure of actophotin from a cloned cDNA, determine the 3-D structure of actophorin at atomic resolution, identify amino acid contacts between actophorin and actin by chemical crosslinking and protein sequencing so that we can show how the proteins bind together, characterize the interaction of actophorin with actin monomers and filaments as a model for the mechanisms of other severning proteins, search for factors that might regulate the interaction of actophorin and actin and localize actophorin in Acanthanoeba. To characterize the actin filament crosslinking protein called alphaactinin, we will study the mechanical properties of mixtures of alphaaclinin and actin filaments as a function of filament length and concentation, analyze the dynamics of the interaction of alphaactinin with actin filaments by fluroescence recovery after photobleaching, stopped flow kinetics, equilibrium binding and electron microscopy, search for factors regulating the interaction of alphaactinin with actin filaments and document the intracellular dynamics of alphaactinin by microinjection of labeled protein and photobleaching recovery.

Actin is found abundantly in all eukaryotic cells, where it plays a key role in contractile processes, and in the maintenance as well as the dynamics of cell shape and cytoplasmic consistency. In attempts to ultimately understand the molecular basis of these actin-based processes, we propose to use high resolution electron microscopy and three-dimensional (3D) image analysis and processing to develop a consistent 3D molecular model of the functionally important actin filament, and to investigate the structural basis of actin's specific interactions with itself and with various contractile, regulatory and cytoskeletal proteins. Using structural, proteinchemical, immunological and microinjection methods, we also propose to explore the role of mutant actins isolated from stably transformed human fibroblasts in neoplasia. Specifically, we propose to proceed as follows: (1) To determine a consistent 3D model of the actin molecule at the 1-2 nm resolution level from different types of negatively stained and frozen hydrated crystalline actin arrays. (2) To develop a consistent 3D molecular model of the actin filament from a variety of negatively stained and frozen hydrated actin filament arrays and to align molecular actin models within the resulting filament 3D reconstructions. (3) To map binding sites for contractile, regulatory and cytoplasmic proteins on the actin molecule via 3D structural analysis of stoichiometrically decorated crystalline actin arrays or from co-crystals induced from specific complexes between actin and actin-interacting proteins. (4) To structurally and functionally analyze several types of crosslinked actin dimers (a) to aid in developing a consistent molecular model of the actin filament (see 2) and (b) to understand their role in regulating actin polymerization. (5) To investigate the in vitro polymerization properties, 3D structure, and in situ distribution (i.e. via immunofluorescence with mutant actin-specific antibody) and interactions (i.e. via microinjection of mutant actin mixtures) of mutant beta-actins isolated from transformed human fibroblasts.

The long range goals of this research program are to (1) establish the structural changes in the actomyosin complex that produce motile force coupled to ATP hydrolysis, (2) characterize the mechanism of organelle movements by myosin-1 and (3) determine the mechanisms that control the accumulation of myosin and its activation in the cleavage furrow during cytokinesis. To reach these goals, we propose the following studies: Project 1, Electron microscopy of intermediate in the actomyosin cycle. We will use a novel rapid-mixing/stopped-flow rapid- freezing device to capture the myosin intermediates that are weakly bound to actin filaments in the actomyosin ATPase cycle. After freeze-fracturing, the structure of these intermediates will be studied directly by electron microscopy to establish the structural changes that produce force and motion. Project 2. Assembly and function of myosin-II. To characterize the molecular events in the assembly and function of myosin filaments in non-muscle cells, we will use a variety of biophysical methods to establish the mechanism of assembly and its regulation by other proteins. This will include the effects of monoclonal antibodies on assembly, ATPase activity and in vitro assays for motility. cDNA's for the molecule will be expressed in bacteria to map precisely the binding sites for more than 40 monoclonal antibodies, to provide antigens for the production of new monoclonal antibodies and to produce myosins modified by deletions or point mutations in vitro for biochemical tests for function. Project 3. Interaction of myosin-1 with membranes. We will carry out quantitative assays to characterize the binding of myosin-1 to the membranes of isolated organelles and test the reconstituted membranes for their ability to move along actin filaments. To establish the membrane binding region of the myosin-1, we will test well defined segments of the protein expressed in bacteria from cloned cDNAs for their ability to bind to membranes. We will also carry out in vitro assays of actin motility with myosin-1 to establish the regulatory functions of 2 classes of heavy chain kinases. Project 4. Regulation of cytokinesis. We will produce monoclonal antibodies to the phosphorylated sites of the myosin light chain and use them to map out the times and places where myosin is phosphorylated during cell division. In parallel experiments we will modify isolated myosin light chains with a fluorescent dye so that after they are microinjected into live cells (where they will recombine with myosin heavy chains) we can follow the movements of the myosin during cell division. By also modifying these fluorescent light chains with thiophosphate on either the protein kinase C site or the myosin light chain kinase site, we will establish whether either or both of these modifications influence the movement of the myosin into the cleavage furrow at cytokinesis.

[unreadable] DESCRIPTION (provided by applicant): Our goal is to understand the molecular mechanisms controlling assembly and disassembly of actin filaments during cellular locomotion and endocytosis in terms of the rates of specific reactions. We will use a combination of biochemical, genetic and cellular experiments in fission yeast to test how cells maintain a pool of actin subunits, initiate new actin filaments and disassemble aged actin filaments. We will use spectroscopic assays to map the pathway of actin filament branch formation by Arp2/3 complex in the presence and absence of profilin. We will characterize the biochemical properties of a temperature sensitive mutant of fission yeast cofilin for use in experiments with live yeast cells. We will reconstitute a recycling actin motility system from purified proteins and analyze the dynamics of individual filaments by fluorescence microscopy. We will use fluorescence microscopy of fission yeast expressing functional fusion proteins to determine the molecular pathway of actin filament assembly and turnover during endocytosis for comparisons with mathematical models. We will test mechanisms in cells by varying protein concentrations and/or mutagenesis. We will use direct observations of growing actin filaments to determine how elongation is influenced by the length of linker peptides in the FH2 domain, tension on the N-terminus of formin FH1FH2 constructs and the distance between profilin binding sites in the FH1 domain and the FH2 domain. We will use single molecule fluorescence microscopy to study conformational changes in the FH2 domain on the end of actin filaments and to investigate the "rotation paradox" of FH2 domains on the ends of growing actin filaments. [unreadable] [unreadable] PUBLIC HEALTH REVELATION: We study the molecular basis of cellular motility and cytokinesis, particularly the roles of actin filaments and myosin motors. Actin-based movements are essential for cell division, shaping organs during embryonic development, defense against microorganisms and wiring the nervous system. Movement of cells out of primary tumors is the chief cause of mortality in cancer. [unreadable] [unreadable] [unreadable]

Our goals are to understand the mechanisms of (1) assembly of myosin-II into bipolar filaments, (2) cytokinesis and (3) organelle movements by myosin-I. To reach these goals, we propose the following studies: 1. Assembly and function of myosin-II. We will determine the atomic structure of the C-terminal part of the tail of Acanthamoeba myosin-II that is required for the formation of minifilaments. We will compare the assembly kinetics of C-terminal fragments with intact myosin tails to test our hypothesis that assembly is favored by intramolecular diffusion. 2. Regulation of cytokinesis. We will use biochemistry, genetics and molecular genetics to study cytokinesis in Schizosaccharomyces pombe. We will determine genetic interactions between the two isoforms of myosin-II and gene products known to participate in cytokinesis. After identifying and characterizing the light chains associated with the two isoforms of myosin-II, we will use genetic screens to identify new components that regulate the time and position of furrow formation. 3. Interaction of myosin-I with membranes. We will characterize lipid binding properties of Acanthamoeba myosin-I isozymes, analyze the assembly of myosin-I tails into higher order structures in solution and on lipid surfaces and determine which parts of the myosin-I molecule are required to target the three isoforms to specific membranes in vivo. To aid in this analysis, we will attempt to crystallize the tail myosin- I for high resolution structure determination. This work will contribute to our understanding of basic cellular process such as cell division (as important feature of cancer), cellular motility (important for tumor metastasis), phagocytosis (important in defense against microbial pathogens) and muscle contraction.

DESCRIPTION (provided by applicant): My long range goal is to understand the mechanism of cytokinesis. We will pursue complementary cellular, biochemical and genetic strategies using the fission yeast Schizosaccharomyces pombe to address the following specific aims: Project 1. Dynamics of contractile ring proteins: these quantitative experiments on molecules in live cells-will provide the parameter values required to decipher the pathway of contractile ring assembly. We will measure the rates that cytokinesis proteins exchange between the cytoplasm and contractile rings, determine if actin filaments influence the dynamics of each contractile ring protein and determine if cytokinesis proteins influence each other's dynamics. Project 2. Assembly of cytokinesis proteins in the broad band of precursor nodes: these experiments will clarify the biochemical pathway that assembles the precursor of the contractile ring. We will determine the domains of anillin Mid1p that bind the cortex and downstream proteins and how cytokinesis proteins interact with Mid1p and with each other. Project 3. Transformation of precursor nodes into a contractile ring: this work will provide information required to understand the second step in the assembly of the contractile ring. We will investigate how Cdc12p controls the assembly of actin filaments from nodes, characterize the mechanical process that converts, a broad band of nodes into a contractile ring, characterize how DCS protein Rng3p and phosphorylation affect the structure of Myo2 filaments and determine how cytokinesis proteins are organized in the broad band and the contractile ring. Project 4. Constriction and disassembly of the contractile ring: these experiments will help us understand how cells trigger constriction of the contractile ring and how the ring disassembles as it constricts. We will investigate how the SIN pathway triggers constriction of the contractile ring, explore the factors that limit the rate of constriction of contractile rings and determine how the contractile ring disassembles during constriction.

DESCRIPTION (provided by applicant): The long-term goal of this program project is to understand the molecular mechanisms that allow Arp2/3 complex to control actin filament assembly during endocytosis and cellular motility. The mechanism of action of Arp2/3 complex is so complicated that a team effort is required. Our group of four geographically dispersed laboratories combines x-ray crystallography, electron microscopy, image processing and molecular biology to address technically challenging questions. Structural insights from his ongoing collaboration will provide important clues about reaction mechanisms and also help to design experiments to measure kinetic and thermodynamic constants and to study actin assembly in live cells. Project 1 (Pollard) proposes a combination of biochemical and x-ray crystallographic experiments to obtain detailed structural information regarding conformational changes and macromolecular interactions of Arp2/3 complex. Project 2 (Li) proposes a combination of biophysical studies on interaction's of nucleation promoting factors with purified Arp2/3 complex and microscopic analysis of mouse cells lacking subunits of Arp2/3 complex. Project 3 (Hanein) proposes electron microscopy of purified Arp2/3 complex, actin filament branches and of actin networks in mouse cells with and without functional Arp2/3 complex. Project 4 (Volkmann) proposes computational methods to extract the maximal amount of information from spectroscopy experiments in projects 1 and 2 and the electron micrographs from project 3.

We will use crystallography and radiation foot printing to learn how nucleotides and activating peptides influence the conformation of Arp2/3 complex. This structural information will contribute to the group effort to model the entire structural pathway of Arp2/3 complex activation and nucleation of actin filament branches. Project 1. We will determine the structures of activator peptides derived from WASp, N-WASp or Scar/WAVE1 bound to Arp2/3 complex by x-ray crystallography. We will use mutation and biochemical characterization to characterize the contributions of key residues of activator proteins for binding Arp2/3 complex. Using insights from NMR and crystallography we will design and characterize biochemically activator proteins with mutations that might compromise their binding to or activation of the Arp2/3 complex. Project 2. Determine how nucleotides and WASp/Scar proteins influence the equilibrium between the active and inactive conformations of Arp2/3 complex. We will obtain high resolution structures of active conformations of Arp2/3 complex to document the conformational changes required to activate the complex for nucleating actin filaments and synchrotron radiation protection to identify residues in Arp2/3 complex that change their exposure to the solvent upon binding nucleotides and activator peptides. Project 3. Determine how Arp2/3 complex initiates an actin filament branch. We will crystallize and determine the structures of actin with a bound WH2 peptide or p40 insert peptides. Use x-ray crystallography to determine the structure of activated Arp2/3 complex with the first subunits of the nucleated actin filament. We will attempt to crystallize Arp2/3 complex associated with WA and one or two actin monomers. Project 4. Determine the structure of actin filament branches mediated by Arp2/3 complex. We will use synchrotron radiation protection to identify amino acid residues at interface between Arp2/3 complex and the side of actin filaments.

intermolecular interaction; fluorescent dye /probe; molecular site; crosslink; biomedical resource;

The assembly dynamics of the actin cytoskeleton is crucial for most morphogenetic processes that occur at both cellular and organism levels in eukaryotes. As a consequence, defects in actin dynamics and organization have been implicated in a variety of human diseases. However, our understanding of how actin is regulated in different physiological processes remains limited due to the extraordinary complexity and dynamic nature of the actin cytoskeletal system. The Arp2/3 complex is thought to be a major key element of actin filament nucleation and the formation of branched dendritic networks at the leading edge of motile cells. The role of Arp2/3 complex in these processes can only be understood through studies of the key molecular steps in the Arp2/3-mediated actin nucleation and branch formation. In this Program Project we aim at understanding the detailed structural mechanisms by which the Arp2/3 complex mediates the formation of actin branch junctions and the role of the Arp2/3 complex in vertebrate cell motility. We have assembled a highly synergistic team of Pis with complimentary expertise to achieve these goals through collaborative efforts. This sub-project (Volkmann lab) will contribute the development and application of a diverse set of computational tools for the reconstruction, analysis, and modeling of structural states and distribution patterns. Using these tools we will combine information from various data sources generated by all members of the Program Project to obtain (i) high-resolution models of the intermediates of the branch formation process and of the fully assembled branch itself;(ii) a dynamic, energetically self-consistent, structural model of the transition pathway from the inactive state of the Arp2/3 complex to the fully assembled branch in the dendritic network, and (iii) structural differences between different cell types and between wild-type cells and defective mutants by multi-dimensional and dynamical characterization of Arp2/3-complex and actin-filament distribution patterns in living eukaryotic cells.

DESCRIPTION (provided by applicant): Dendritic spines and their associated synapses become prematurely destabilized in psychiatric and neurodegenerative diseases. Proper control of the actin cytoskeleton is critical for the long-term structural stability of dendritic spines, bt currently little is known about the molecules and mechanisms that confer long-term structural stability on spines and the field remains understudied. We discovered that loss of integrin ?3?1 signaling through the Abl2/Arg nonreceptor tyrosine kinase causes widespread dendrite arbor loss and dendritic spine destabilization. Even though Arg inhibits the RhoA GTPase to stabilize dendrite arbors, this mechanism does not impact spine stability, raising the fundamental question of how Arg stabilizes spines. We provide evidence that Arg directly binds and stabilizes actin filaments and also regulates the binding and actions of the actin regulators cortactin and Arp2/3 complex on actin filaments. We also find that Arg-mediated recruitment of cortactin to dendritic spines is crucial for spine stability. Our proposal will test the highly innovative hypothesis that Arg interacts physically and functionally with actin filaments and actin regulatory proteins to directly regulate actin dynamics and thereby stabilize dendritic spines. Our first aim will elucidate how Arg:cortactin interactions control actin dynamics. We find that Arg binding to actin filaments stabilizes them from depolymerization. Arg binding also recruits the actin-binding protein cortactin, which stabilizes actin filaments and increases actin branch formation by Arp2/3 complex. We will use total internal reflection microscopy to observe single filaments and to measure how Arg and cortactin affect actin filament stability, Arp2/3 complex-mediated branch formation, and cofilin-mediated actin filament severing. We will use mutants of these proteins that do not interact with each other or with actin filaments to identify which protein:protein interaction interfaces are critical for effects on actin dynamics. These studies will reveal how Ar and cortactin affect actin filament stability, branching, and turnover. Our second aim will determine how Arg and cortactin modulate spine stability via effects on actin dynamics. We find that knockdown of Arg in neurons results in the loss of cortactin from spines and triggers their destabilization. We hypothesize this destabilization is due to the disruption of normal actin dynamics in spines. Knockdown of Arg or cortactin in established hippocampal neuron cultures compromises dendritic spine stability. These deficits can be quantitatively rescued by re-expression of shRNA-resistant versions of Arg or cortactin, respectively. Employing our collection of Arg and cortactin mutants, we will test how mutational disruption of key interaction interfaces in these proteins affects dendritic spine shape and stability. We will use fluorescence recovery after photobleaching (FRAP) of GFP-actin in spines to reveal how manipulations of Arg and cortactin function affect actin dynamics in spines and determine how this relates to the effects of these proteins on actin biochemistry and spine stability.

? DESCRIPTION (provided by applicant): The long-range goal of the project is to understand the mechanism of cytokinesis in enough detail to make useful mathematical models of the process that can predict the results of future experiments. Remarkably, we are close to this goal for the fission yeast S. pombe owing to the experimental advantages of this organism. Our 2008 model for contractile ring assembly from cytokinesis organizing centers called nodes faithfully accounts for prior experimental observations and our subsequent work. Over the past 4 years we tested new models for the formation of cytokinesis nodes from two types of interphase nodes and for the constriction and turnover of the contractile ring. Simulations of these three models show that we have a good understanding of the physical events relating to cytokinesis around the entire cell cycle. The constriction model generates the tension observed in live cells and explains why the constant turnover of both actin filaments and myosin is required for constriction. This work puts us in position to ask well-informed questions about the mechanisms that control each of the transitions in the process. Our first goal for the next award period is to determine the structure of nodes, the cytokinesis-organizing centers of fission yeast. We aim to determine how ten different proteins are organized in interphase and cytokinesis nodes, including node protein Blt1p, exchange factor Gef2p, cell cycle kinases Cdr1p and Cdr2p, anillin Mid1p, two myosin-II isoforms, F-BAR Cdc15p, formin Cdc12p and IQ-GAP Rng2p. We will combine information from (i) biochemical and biophysical characterization of each protein, (ii) SAXS and x-ray crystallography of selected protein domains and (iii) super-resolution fluorescence microscopy of live cells. The second goal is to characterize the life cycles of the two types of interphase nodes and their combination to form cytokinesis nodes. Observations of cells with mutations in regulatory proteins will reveal how the cell cycle controls the transitionsin the node cycle such as the disappearance of type 1 nodes during mitosis. The third goal is to use super-resolution microscopy of live cells, modeling and effects of mutations to characterize the dynamics of the protein components of the contractile ring as it constricts and disassembles. These projects are powered by four technical innovations. (1) Our method to count fluorescent molecules in confocal images is the basis of our quantitative approach to microscopy of live cells. (2) We have taken quantitative microscopy to a new level with a novel method to measure affinities in live cells. (3) A superior photoswitchable protein and high-speed image acquisition allowed us to make real time super-resolution microscopy routine for live fission yeast. (4) We expanded our mathematical models of cytokinesis to include the formation of cytokinesis nodes from two types of interphase nodes and constriction of the contractile ring. Given the evolutionary conservation of many of the participating molecules, I believe that studies of fission yeast will establish the basic molecular pathways controlling cytokinesis in other eukaryotes.