John Heuser - US grants

The quick-freeze, deep-etch technique developed in this laboratory will be used to study three key processes in eukaryotes, namely, the formation and dissolution of clathrin cages, the assembly and disassembly of an extracellular matrix, and recognition between gametic cells. Ongoing studies of all three systems are described to demonstrate the unique contributions the quick-freeze, deep-etch technique can make to their understanding. Critical is the ability of the technique to visualize proteins in their native state and after experimental manipulation. Thus, for example, the components of the clathrin cages can be viewed as individual proteins ("triskelions", "cagin", etc.) or in various stages of assembly, disassembly, and association with membrane surfaces. Particularly valuable are images of interactions between fibrous proteins, such as those that constitute the extracellular matrix and the sexual agglutinins of Chlamydomonas, since these are impossible to decipher by any other electron microscopic technique. A broad range of experiments is proposed to probe the "rules" by which these self-assembling sets of proteins carry out their biological functions. Many of these experiments will involve the structural analysis of proteins purified in two collaborating laboratories; in addition, new approaches are described for studying such processes as clathrin/membrane interactions. These studies will hopefully increase our basic understanding of how extracellular proteins interact, how cells regulate their uptake of proteins and their internal membrane traffic, and how cells recognize each other.

This project will continue to develop new techniques and machinery for rapidly freezing skeletal muscles as they are stimulated and monitored electrophysiologically. Neuromuscular junctions frozen at the precise moment of neurotransmission will be prepared by the methods of freeze fracture and freeze substitution for examination in the electron microscope. In this way, the fleeting structural changes that occur during secretion of acetylcholine from the nerve and its reception by the muscle will be revealed in their natural form. This structural data will help to elucidate the molecular mechanisms that underlie normal neuromuscular transmission, and will form a basis of comparison with neuromuscular pathology. Work will aso continue on the new quick-freeze, freeze-dry technique which promises to elucidate the detailed interrelationships between membrane molecules and cytoplasmic filaments in a variety of cellular processes, including hormone uptake, organelle translocation, and nonmuscle cell motility.

The freeze-etch electron microscopic techniques developed in this laboratory will be used to study clathrin-mediated endocytosis in fibroblasts and macrophages. Questions to be addressed include: (1) what role do clathrin lattices play in bringing about receptor clustering? (2) what is the mechanism by which clathrin lattices curve into spheres and pinch off from the plasma membrane? and (3) how is clathrin recycled from internalized membranes back to the cell surface for additional round of endocytosis? Ongoing studies of all three processes are described to demonstrate the unique contributions that the quick-freeze, deep-etch, rotary=replication technique can make to this analysis. By permitting direct visualization of individual membrane receptors on the outsides of cells, as well as individual clathrin lattices on their insides, the technique has already demonstrated a number of correlations between clathrin lattice dynamics and changes in receptor distribution. By also permitting 3-D visualization of small clathrin polymers and individual clathrin "triskelia" adsorbed to mica, the technique has also begun to shed light on the molecular interactions that underlie clathrin lattice dynamics. A broad range of experiments is proposed to further probe the rules by which clathrin molecules self-assemble and carry out their biological functions. Some of these experiments will involve electron microscopic analysis of the effects of chemical treatments that stimulate or inhibit endocytosis in cultured cells, particularly the inhibitory effects of cytoplasmic acidification and the stimulatory effects of cytoplasmic alkalinization. In addition, new observations on the effects of such pH changes on endosome and lysosome movement and fusion will be actively pursued. Finally, the molecular basis of clathrin lattice dynamics will be analyzed by observing the patterns of molecular interaction that clathrin displays with itself and with other proteins in vitro, under different conditions that mimic the above in vivo situations. These studies will hopefully increase our basic understanding of how cells regulate their uptake of proteins and their intracellular membrane traffic.

The freeze-etch electron microscopic techniques developed in this laboratory will be used to study clathrin-mediated endocytosis in fibroblasts and macrophages. Questions to be addressed include: (1) what role do clathrin lattices play in bringing about receptor clustering? (2) what is the mechanism by which clathrin lattices curve into spheres and pinch off from the plasma membrane? and (3) how is clathrin recycled from internalized membranes back to the cell surface for additional round of endocytosis? Ongoing studies of all three processes are described to demonstrate the unique contributions that the quick-freeze, deep-etch, rotary=replication technique can make to this analysis. By permitting direct visualization of individual membrane receptors on the outsides of cells, as well as individual clathrin lattices on their insides, the technique has already demonstrated a number of correlations between clathrin lattice dynamics and changes in receptor distribution. By also permitting 3-D visualization of small clathrin polymers and individual clathrin "triskelia" adsorbed to mica, the technique has also begun to shed light on the molecular interactions that underlie clathrin lattice dynamics. A broad range of experiments is proposed to further probe the rules by which clathrin molecules self-assemble and carry out their biological functions. Some of these experiments will involve electron microscopic analysis of the effects of chemical treatments that stimulate or inhibit endocytosis in cultured cells, particularly the inhibitory effects of cytoplasmic acidification and the stimulatory effects of cytoplasmic alkalinization. In addition, new observations on the effects of such pH changes on endosome and lysosome movement and fusion will be actively pursued. Finally, the molecular basis of clathrin lattice dynamics will be analyzed by observing the patterns of molecular interaction that clathrin displays with itself and with other proteins in vitro, under different conditions that mimic the above in vivo situations. These studies will hopefully increase our basic understanding of how cells regulate their uptake of proteins and their intracellular membrane traffic.

The goal of this project is to systematically study the structural underpinnings of exocytosis and endocytosis in a relevant and tractable cell-culture system that properly reflects the special properties of neurons and neuroendocrine cells. This reflects the applicant's long-term interest in elucidating the mechanisms of membrane recycling that link exo-and endocytosis at the synapse, where these mechanisms are fundamental to neuronal communication, learning, and memory. The applicant has chosen to study PC12 cells (derived from an immortalized rat pheochromocytoma) because they have been well-characterized biochemically and physiologically, they have a known complement of neuronal-type proteins involved in both secretion and endocytosis, and they have been used extensively in the field, to date. The applicant's specific approach will be to apply the special technique of "deep-etch" electron microscopy (EM) pioneered in his laboratory to obtain 3-D images of structural changes that occur in PC12 cells attached to adhesive substrates and "unroofed" with an ultrasonic jet of buffer to expose their inner membrane surfaces. That is where all the exo-and endocytotic events occur, and where the applicant intends to witness membrane changes and identify their molecular underpinnings. To accomplish this goal, the applicant will use his lab's new developments in digital/computer-based techniques for creating "anaglyph" 3-D images of "deep-etch" EM's, combined with novel methods of identifying molecules and organelles by :1) EM immunocytochemistry with gold-labeled secondary antibodies and 2) transiently expressed epitope-tagged molecules that can either be seen directly by "deep etch" EM, or secondarily after DAB-based histochemistry is performed on them. This will permit the applicant to establish a close correlation between the "static" images of PC12 cells obtained by EM and the "dynamic" images of PC12 cells they plan to obtain by real-time confocal light microscopy. By this combination of powerful and modern microscopic imaging techniques, the applicant hopes to deepen our understanding of the basic cellular mechanisms that mediate neuronal communication, which should help to advance neuropharmacology and therapeutics and begin to address some of the fundamental questions about what goes wrong with synaptic communication in neuronal degenerative diseases such as Alzheimer's.

DESCRIPTION (provided by applicant): Our procedures for "deep-etch" EM will be modified in the coming period to include "high-pressure slam-freezing", so that we can obtain new structural data on the molecular events that occur within the cytoplasm and membranes of virally-infected and virally-transformed whole cells. This will involve, first, completing the development of a new "high pressure slammer" that we have invented (and making it available to the research community at large). Second, we will apply this new technology to the particularly vexing and challenging problem of how poxviruses are manufactured and disseminated by animal cells. Here we will use attenuated strains of Vaccinia virus (VV) that can be handled completely safely, to be provided by Dr. Bernard Moss of the NIH, who has generated a whole host of mutant VV's that will allow us to determine what phenotypic variations these mutations cause at the EM level, and thus to better understand the molecular mechanisms behind poxvirus infection. Third, we will exploit our new freezer's capacity to capture rapid membrane changes by imaging the dynamic "invadopodia" of one type of cancer cell, the RSV-transformed BHK cell, which forms unique "podosome rosettes" on its ventral surface that will serve as ideal 'test patterns' for us to identify transient physical changes in the organization of proteins and lipids (especially "signaling rafts") the cell membrane, and to determine what membrane shape-changes are associated with these phase-dynamics. This and our next project will benefit from a further technical advance we have made, which permits us to freeze cells right through their coverslips and thus optimally image their ventral surfaces in the EM. In close work with Dr. Phyllis Hanson, who has developed a number of mutant ESCRT's and mutant AAA-ATPases that perturb MVB formation, we have used this technique to show that certain of her mutants redirect MVB budding to the cell surface, where we can easily 'capture' it in the EM. This promises to shed much light on the molecular mechanisms involved in various sorts of membrane budding, and to reveal the remarkable parallels between processes as seemingly different as multivesicular body formation and the budding of retro viruses from the cell surface.

[unreadable] DESCRIPTION (provided by applicant): This application requests the funds to purchase a new, digital electron microscope to replace one of the two 30-year-old, nearly-obsolete electron microscopes that constitute the unique "deep-etch" EM facility that operates at Washington University School of Medicine. This special form of electron microscopy has long been recognized as a uniquely valuable tool for biomedical research, and is a national resource that this institution wishes to promote and support long into the future. The deep-etch technique permits the visualization of all sorts of medically-relevant samples, from whole tissues down to individual cells and molecules, in all cases by quick-freeze and freeze-etch procedures that optimally preserve a lifelike state and avoid the artifacts inherent in most other forms of electron microscopy. It yields dramatic three- dimensional images of membranes and membrane topology that have provided critical insights into a wide range of biological problems, from how nerves communicate with each other, to how cancer cells migrate and invade tissues, to how viruses infect cells and replicate inside of them, to mention just a few topics. Deep-etch electron microscopy is thriving, and likely will be further developed and improved with time. However, the workhorse electron microscopes that are currently being used to do it are seriously out-of- date, and by being entirely film-based, have become serious bottlenecks in the process. With a modern electron microscope that includes a top-notch digital camera, the Deep-Etch EM Facility at Washington University School of Medicine will be able to greatly increase its output and streamline its operations, better train the next generation of electron microscopists, and support a wider range of research projects. Thus, this application is a request for a contemporary electron microscope to modernize an existing facility that is already of great value to Washington University and to all of medical science, but which is operating with antiquated equipment. [unreadable] [unreadable] [unreadable]