Dozens of proteins control the docking and fusion of exocytic vesicles in neurons and endocrine cells. The identity and roles of many of these proteins have been assigned through a combination of genetics, biochemistry, and electrophysiology. However, the spatial organization, heterogeneity, regulation, and dynamics of these proteins have yet to be determined. Finding this organization is key to understanding how proteins regulate these systems in healthy cells and might malfunction in disease. Thus, we aimed to map key proteins proposed to act during exocytosis. To accomplish this, we developed a combination of high-throughput live cell imaging, super-resolution fluorescence imaging, and electron microscopy. Through this multi-modal imaging approach the location, dynamics, and occupancy of individual proteins were mapped at specific populations of vesicles in cells and compared to the underlying cellular architecture that organizes the plasma membrane. This allowed us to determine the fundamental organization of the exocytic membrane system and how specific molecular components responsible for vesicle trafficking and fusion assemble together and function in time and space. Specifically, with TIRF microscopy and high-throughput image analysis we developed a universal map of the proteins that control exocytosis and provide a global network level analysis of vesicle fusion events. We were able to identify unique classes of key regulatory molecules that strongly associate with the vast majority of exocytic vesicles in cultured neuorendocrine PC12 chromaffin cells. Specific proteins we identified were Rabs and Rab effectors, SNARE proteins, SNARE modulators, and BAR-domain proteins, and mechanoenzymes including Dynamin. To determine the local dynamics of these components at single sites of exocytosis we imaged local changes of proteins at the exact moment of fusion in live cells. In these studies we discovered an unexpected recruitment of several important endocytic proteins to sites of synaptic-like microvesicle fusion. These molecules include the regulatory proteins dynamin, amphiphysin, syndapin, and endophilin. We further discovered that mutations of several of these proteins altered the kinetics of vesicle membrane protein release into the plasma membrane. Our hypothesis is that specific proteins (dynamin, syndapin, amphiphysin, and endophilin) regulate the dilation or permeability of the fusion pore to control the amount of membrane-bound cargo released during single exocytic fusion events. This would allow excitable cells to modulate the amount of material released during even single fusion events. In an effort to develop new and better imaging probes for tracking exocytosis of vesicles in living cells, we collaborated with two other labs to build and test new organic red-fluorescence pH sensitive imaging probes. Working together with these groups (a chemistry group and neurophysiology group), we synthesized and tested a semi-synthetic pH-sensitive red fluorophore for tracking exocytosis and endocytosis in cultured neuroendocrine cells and neurons. We were able to show that this dye/protein system performed as good as the best available green fluorescent protein-based pH sensors for tracking exocytosis. These new probes allow for the high-contrast monitoring of different populations of cells or vesicles in multiple colors and deeper into tissues. We discovered that in cells after exocytosis vesicle material is captured on a dense network of pre-formed clathrin-coated structures following exocytosis. Despite the identification of many molecular components of clathrin-mediated endocytosis, a structural understanding of how these molecules come together to build and retrieve material from the plasma membrane during endocytosis is incomplete. In this aim we sought to directly determine the structure of clathrin-coated vesicles responsible for endocytosis in mammalian cells. By understanding how proteins that have been functionally implicated in endocytosis assemble together at the nanoscale we can place decades of biochemistry, cell biology, and genetics into a physical model of membrane retrieval. Dozens of proteins capture, polymerize, and reshape the clathrin lattice during clathrin-mediated endocytosis (CME). How or if this ensemble of proteins is organized in relation to the clathrin coat is unknown. To determine this nanoscale structure, we developed a super-resolution correlative light and electron microscopy imaging method. This allows us to image the nanometer-scale location of proteins in the context of their local cellular environment. Specifically, we succeeded in developing a robust pipeline for imaging the plasma membrane of cells with 2D localization microscopy and transmission electron microscopy (TEM) of platinum replicas. In these studies we imaged the nanoscale position of endocytic proteins at single clathrin-coated structures. We localized 19 other endocytic proteins (amphiphysin1, AP2, 2-arrestin, CALM, clathrin, DAB2, dynamin2, EPS15, epsin1, epsin2, FCHO2, HIP1R, intersectin, NECAP, SNX9, stonin2, syndapin2, transferrin receptor, VAMP2) on thousands of individual clathrin structures, generating a comprehensive molecular architecture of endocytosis with nano-precision in human Hela cells. From this work, we discovered that endocytic proteins distribute into distinct spatial zones (rings) in relation to the edge of the clathrin lattice. The presence or concentrations of proteins within these rings change at distinct stages of organelle development. We propose that endocytosis is driven by the recruitment, reorganization, and loss of proteins within these partitioned nano-scale zones. In total, these studies are allowing us to build structural models for how proteins are organized at single organelles to regulate endocytosis, a central process for all eukaryotic cells. Clathrin-coated pits dynamically assemble and disassemble at the membrane of mammalian cells. Models of how clathrin coats curve have been controversial. Specially, some models propose that clathrin can only grow as a curved lattice while others propose that clathrin first assembles as a fully formed flat lattice that later curves into a sphere. We tested these models in two collaborative projects by imaging growing clathrin structures with both live cell polarized total internal reflection microscopy (pTIRF) and super-resolution EM correlative imaging. In these studies we watched single pits curve at the plasma membrane and related these observations to structures observed in EM. Our work demonstrated that the pathway for curvature is heterogenous. First, many clathrin sites form as small curved structures, others grow partially flat and then begin to curve, and some grow to their full size and bend. Thus, cells control the pathway of membrane curvature with multiple mechanisms of curvature generation in the lattice. Because of the above findings, it was clear that even in a single cell the pathway of clathrin assembly is mixed. To test if specific cell types have unique clathrin assembly pathways we again collaborated to explore how the endocytic system changed when stem cells were differentiated into unique states including neural progenitor cells and fibroblasts. Surprisingly, we discovered that even in isogenic stem cells, the clathrin system at the plasma membrane was radically changed after these cells were differentiated. Specifically, stem cells and NPCs had small highly-active clathrin coated pits while fibroblasts grew very large, domed, clathrin structures that were much slower and more stationary. We hypothesize that cells tune their endocytic system according to their unique biophysical, metabolic, and signaling needs of that system.
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