DYNAMICS OF SECRETORY MEMBRANE TRAFFICKING, SORTING, AND COMPARTMENTALIZATION
, Head, Section on Organelle Biology
George Patterson, PhD, Staff Scientist
Jennifer Chua, PhD, Postdoctoral Fellow
Jennifer Gillette, PhD, Postdoctoral Fellow
Peter Kim, PhD, Postdoctoral Fellow
Wei Liu, PhD, Postdoctoral Fellow
Holger Lorenz, PhD, Postdoctoral Fellow
Manos Mavrakis, PhD, Postdoctoral Fellow
Kasturi Mitre, PhD, Postdoctoral Fellow
Carolyn Ott, PhD, Postdoctoral Fellow
Richa Richy, PhD, Postdoctoral Fellow
Rachid Sougrat, PhD, Postdoctoral Fellow
Christian Wunder, PhD, Postdoctoral Fellow
Dale Hailey, BA, Predoctoral Fellow
Development of green fluorescent protein technology
Over the past year, we have continued to develop new fluorescence imaging approaches, particularly a high-resolution microscopy technique capable of optical resolutions beyond the limit imposed by diffraction. We developed the technique in collaboration with Eric Betzig, Harald Hess, and members of Juan Bonifacino’s and Michael Davidson’s laboratory (Betzig et al., Science 2006;313:1642). Termed photoactivated localization microscopy (PALM), the method involves serial photoactivation and subsequent bleaching of several sparse subsets of photoactivated fluorescent protein molecules. PALM localizes individual molecules at near-molecular resolution by determining their centers of fluorescent emission via a statistical fit of their point-spread function. It then assembles the aggregate position information from all subsets into a super-resolution image that isolates individual fluorescent molecules at high molecular densities (up to 105 molecules/µm2). We demonstrated PALM imaging of intracellular structures (including lysosomes, Golgi apparatus, and mitochondria) in cryo-prepared thin sections, vinculin and actin in fixed cells with TIRF (total internal reflection) excitation, and correlative PALM/transmission electron microscopy of a mitochondrial marker protein. We also worked toward developing dual-label PALM by using two different photactivatable molecules expressed within cells. In addition, we developed a system for single-particle tracking by using PALM in living cells, allowing characterization of protein diffusion and immobilization at the single-molecule level.
A second new fluorescent protein technique—fluorescence protease protection (FPP)—developed in our laboratory permits determination of a protein’s topology in living cells (Lorentz et al., Nat Methods 2006;3:205; Lorentz et al., Nat Protocols 2006;1:276). The assay provides a fluorescent readout in response to trypsin-induced destruction of GFP attached to a protein of interest before and after plasma membrane permeabilization. In performing the FPP assay, we attach a fluorescent protein to the N- or C-terminus of a protein. Subsequently, cells expressing the fusion protein are exposed to trypsin either before or after plasma membrane permeabilization by digitonin. If the fluorescent protein moiety on the expressed protein faces the environment exposed to trypsin (that is, the cytoplasm), then its fluorescent signal is lost. Conversely, if the fluorescent protein moiety on the expressed protein faces the environment that is protected from trypsin (that is, the lumen of a compartment), then its fluorescence persists. Given these outcomes and the fluorescent protein’s known engineered position within the protein, it is possible to deduce the orientation of the protein across the lipid bilayer. We demonstrated the broad applicability of FPP by using it to define the topology of proteins localized to several organelles, including the ER, Golgi apparatus, mitochondria, peroxisomes, and autophagosomes.
Lippincott-Schwartz J. Anything can work. Biotechniques 2006;40:19.
Lippincott-Schwartz J. Biofluorescence: the making of a new technology. Nat Cell Biol 2006;8:1212.
Mavrakis EM, Rikhy R, Lilly MA, Lippincott-Schwartz J. Fluorescence imaging techniques for studying Drosophila development. Curr Protoc Cell Biol 2007, in press.
Wakabayashi Y, Chua J, Larkin JM, Lippincott-Schwartz J, Arias IM. Four-dimensional imaging of filter-grown polarized epithelial cells. Histochem Cell Biol 2007;127:463-72.
Ward TH, Lippincott-Schwartz J. The uses of green fluorescent protein in mammalian cells. Methods Biochem Anal 2006;47:305-37.
Golgi biogenesis and inheritance
While Golgi inheritance during mammalian cell division is known to occur through the disassembly, partitioning, and reassembly of Golgi membranes, the mechanisms responsible for these processes are poorly understood. To investigate such mechanisms, we examined the identity and behavior of Golgi proteins within mitotic membranes by using dynamic cell imaging of Golgi and endoplasmic reticulum (ER) markers, electron microscopy, ER fragmentation with ionomycin, and ER entrapment through misfolding. We drew two overall conclusions from the data. First, the mitotic Golgi haze, seen in metaphase, represents recycled Golgi proteins trapped in the ER, a consequence likely related to the mitosis-specific disassembly of ER exit sites and inactivation of Arf1. Second, mitotic Golgi fragments, seen in prometaphase and telophase, are not isolated breakdown products of the Golgi; rather, they are structures undergoing continuous exchange of their components through the ER and dispersed ER exit sites. These conclusions suggest a model wherein the Golgi is inherited through the ER in mitosis and mitotic Golgi disassembly/reassembly involves the inhibition and subsequent reactivation of cellular activities that control recycling of Golgi components into and out of the ER.
Support for the first of the two conclusions, namely, that the Golgi haze in metaphase cells represents Golgi proteins within the ER, derives from three lines of evidence: (1) the mitotic haze may be resolved into ER by high-resolution confocal microscopy, (2) the haze redistributes with ER into fragments upon ionomycin treatment, and (3) the haze displays quality control features characteristic of ER such as misfolding and retention of proteins. Evidence that mitotic Golgi fragments observed in prophase and telophase represent ER-derived structures through which Golgi proteins cycle rapidly (i.e., from ER exit sites to a fragment and then back again into the ER) came from fluorescence double-labeling, immunoelectron microscopy, and photobleaching recovery experiments. Live-cell imaging of a single cell co-expressing Sec13-YFP and GalT-CFP revealed that mitotic Golgi fragments grow out from ER export domains at the end of mitosis, remain near these sites for a short period, and then undergo clustering into a Golgi ribbon. Immunoperoxidase electron microscopy of cells in prometaphase and telophase further showed that mitotic Golgi fragments were clusters of tubules/vesicles localized adjacent to ER export sites and, in some cases, were in direct continuity with ER export domains. Finally, when a mitotic Golgi fragment was photobleached in cells expressing GalT-YFP, fragment fluorescence rapidly recovered most of the fragment’s original intensity (within two minutes), indicating that Golgi proteins continuously move in and out of mitotic fragments while maintaining steady-state pools in these fragments.
Based on these data, we proposed a multistep process for Golgi inheritance from the ER. In the first step, the formation of prometaphase Golgi fragments near ER exit sites results from (1) accelerated retrograde traffic into the ER and (2) the inability of newly emerging membrane at ER exit sites to recruit the machinery necessary for conversion into Golgi. The next step—appearance of the mitotic Golgi haze—occurs when ER export domains undergo complete disassembly, leading to entrapment of Golgi components in the ER. The final step—reappearance of mitotic Golgi fragments—occurs when ER export domains reassemble and the exported membrane becomes capable of recruiting the machinery necessary for transforming the membrane into the Golgi apparatus. Golgi components exit the ER at ER exit sites and assemble into membrane tubular clusters near these sites. The clusters later transform into Golgi cisternae/stacks and then coalesce into a juxtanuclear site.
Altan-Bonnet N, Sougrat R, Snapp EL, Ward T, Lippincott-Schwartz J. Golgi inheritance in mammalian cells is mediated through ER export activities. Mol Biol Cell 2006;17:990-1005.
ArfGAP1 dynamics and its role in COPI coat assembly
Secretory protein trafficking relies on the COPI coat, which, by assembling into a lattice on Golgi membranes, concentrates cargo at specific sites and deforms the membranes at these sites into coated buds and carriers. The GTPase-activating protein (GAP) responsible for catalyzing Arf1 GTP hydrolysis is an important part of the trafficking system, but the mechanism whereby ArfGAP is recruited to and remains stable within the coat, and what role it plays in maintaining the coat remain unclear. We therefore studied—under different conditions known to affect coat assembly and disassembly—the dynamics of ArfGAP1 by monitoring the membrane turnover of GFP-tagged ArfGAP1 and the GFP-tagged versions of Arf1, coatomer, and ArfGAP1 lacking its zinc finger–containing GAP domain.
We found that all coat components underwent rapid exchange between membranes and cytosol. Focusing on the recruitment process, we discovered that ArfGAP1 binding to Golgi membranes was not dependent on interactions with Arf1 or coatomer, as ArfGAP1∆64N-YFP showed the same Golgi membrane binding properties as ArfGAP1-YFP in photobleaching assays. Moreover, ArfGAP1 recruitment occurred at 4°C, a temperature at which vesicle production is inhibited, thus arguing against a model in which ArfGAP1 targets directly from the cytoplasm onto coated vesicles and then releases. We also found that ArfGAP1 was required to associate with coatomer before it could interact with Arf1 on membranes, as demonstrated by the findings that (1) COPI and ArfGAP1 interacted in biochemical assays and (2) no formation of stable complexes of fluoride-Arf1-ArfGAP1 occurred in ldlF cells depleted of coatomer and treated with AlF. One role for the interaction could be to localize Arf1-ArfGAP1-coatomer complexes spatially on membranes so that ArfGAP1 catalytic activity is confined to Arf1 molecules in association with the COPI coat.
Based on these and previous findings, we proposed a model for ArfGAP1 dynamics in which Arf1 hydrolyzes its GTP within the coat lattice on a time scale that is much more rapid than vesicle budding. The process results in a metastable coat such that the peripheral components constituting the coat undergo constant recruitment to and release from the coat whether or not the membrane buds off as a coated vesicle.
Lippincott-Schwartz J, Liu W. Insights into COPI coat assembly and function in living cells. Trends Cell Biol 2006;10:1-4.
Peroxisome biogenesis
Peroxisomes play an important role in cellular metabolism; they oxidize fatty acids, bile salts, and cholesterol and convert hydrogen peroxide to nontoxic forms, but the locus of peroxisome origination remains unclear. To investigate whether the ER participates in peroxisome biogenesis, we analyzed, in mammalian cells, the trafficking pathway of PEX16, an early-event membrane peroxin. Several results suggested that PEX16 begins life in the ER rather than on peroxisomal membranes. First, biochemical assays revealed that PEX16 incorporates cotranslationally into ER-derived microsomes. Second, PEX16 was localized to the ER in PEX19 mutant cells, which lack peroxisomes. Third, when forced to insert cotranslationally into the ER of live cells through the attachment of a signal sequence, PEX16 transited from the ER to peroxisomes. Finally, upon generation of a variant of PEX16 tagged with PAGFP, we showed that, when the ER pool of PEX16-PAGFP was photoactivated and followed over time, the photoactivated molecules redistributed to peroxisomes.
In experiments employing a photolabeling, pulse-chase strategy for distinguishing newly synthesized from previously synthesized peroxisomal protein components and for visualizing both old and new peroxisomes, we also found that peroxisomal membranes themselves originate from the ER. We discovered that old peroxisomes contained both newly synthesized and previously synthesized protein components, whereas new peroxisomes contained only newly synthesized peroxisomal protein components. This finding argued against fission as the predominant mechanism for mammalian peroxisome formation and indicated that de novo biogenesis of peroxisomes from the ER was important for maintenance of peroxisomes under normal conditions.
Kim PK, Mullen RT, Schumann U, Lippincott-Schwartz J. The origin and maintenance of mammalian peroxisomes involves a de novo PEX16-dependent pathway from the ER. J Cell Biol 2006;173:521-32.
Compartmentalization of the secretory membrane system in the absence of plasma membrane boundaries in the Drosophila syncytial blastoderm embryo
The Drosophila embryo provides a unique model system for studying changes in the organization and dynamics of endomembranes during embryogenesis. Development in the embryo begins with 13 nuclear division cycles within a syncytium and produces more than 6,000 nuclei that, during the next division cycle, become encased in plasma membrane in a process known as cellularization. How the ER and secretory membrane system become equally apportioned among the thousands of synctial nuclei in preparation for cellularization is a question that remains unanswered. To investigate this question in the living embryo, we employed transgenic lines expressing GFP-tagged markers for the ER, Golgi, and plasma membrane in order to examine the organization and morphological changes in the ER, Golgi, and plasma membrane during the precellularization and cellularization stages (Frescas et al., 2006). We found that, before nuclear migration to the embryo cortex, the ER exists as a single, interconnected system through which proteins freely diffuse. Upon nuclear arrival at the cortex, the ER and Golgi become compartmentalized around individual nuclei, as demonstrated in FLIP experiments in which resident proteins of the ER and Golgi rapidly circulate only within the ER and Golgi membrane associated with a particular nucleus during the four rounds of nuclear division at the periphery. Microtubules were necessary for the compartmentalized behavior of the secretory membranes: injection into the embryo of nocodazole to disrupt microtubules caused the ER and Golgi components to circulate more widely in the embryo with no compartmentalization around individual nuclei. These results indicated that, in the absence of plasma membrane boundaries surrounding nuclei but with the requirement of an intact microtubule network, the embryo is able to differentiate the secretory endomembrane system into segregated nuclear-associated units. In a volume occupied by thousands of nuclei, the ability of the embryo to apportion ER and Golgi among nuclei is likely to be vital for both cellularization and the establishment and maintenance of localized gene and protein expression patterns.
Frescas D, Mavrakis M, DeLotto R, Lorenz H, Lippincott-Schwartz J. ER and Golgi membranes exist as restricted and compartmentalized units that surround individual nuclei in the Drosophila syncytial blastoderm embryo. J Cell Biol 2006;173:219-30.
Publications Related to Other Work
Below GA, Altan-Bonnet N, Lippincott-Schwartz J, Ehrenfeld E. Hijacking components of the cellular secretory pathway for replication of Poliovirus RNA. J Virol 2007;81:558-67.
DeLotto R, Steward R, Lippincott-Schwartz J. Nucleocytoplasmic shuttling and dynamic maintenance of the Dorsal/NF-kappaB gradient. Development 2007;134:4233-41.
Mavrakis M, Lippincott-Schwartz J, Stratakis CA, Bossis I. mTOR kinase and the regulatory subunit of protein kinase A (PRKAR1A) spatially and functionally interact during autophagosome maturation. Autophagy 2007;3:151-3.
Mavrakis M, Lippincott-Schwartz J, Stratakis CA, Bossis I. The type IA regulator subunit (RIa) of protein kinase A (PRKAR1A) and mTOR kinase are involved in a common pathway regulating mammalian autophagy. Hum Mol Genet 2006;15:2962-71.
Rey O, Papazyan R, Waldron RT, Young SH, Lippincott-Schwartz J, Jacamo R, Rozengurt E. The nuclear import of protein kinase D3 requires its catalytic activity. J Biol Chem 2006;281:5149-57.
Rossman JS, Stoicheva NG, Langel FD, Patterson GH, Lippincott-Schwartz J, Schaefer BC. POLKADOTS are foci of functional interactions in T-Cell receptor-mediated signaling to NF-kappaB. Mol Biol Cell 2006;17:2166-76.
Snapp EL, Sharma A, Lippincott-Schwartz J, Hegde RS. Monitoring chaperone engagement of substrates in the endoplasmic reticulum of live cells. Proc Natl Acad Sci USA 2006;103:6536-41.
Young ARJ, Chan EY, Hu XW, Kochl R, Crawshaw AR, High S, Hailey DW, Lippincott-Schwartz J, Tooze SJ. Starvation and ULK1-dependent cycling of mammalian Atg8 between TGN and endosomes. J Cell Sci, in press.
1 Nihal Altan-Bonnet, PhD, former Visiting Fellow2 Theresa Ward, PhD, former Visiting Fellow
3 Dave Frescas, BA, former Postbaccalaureate Fellow
COLLABORATORS
Win Arias, MD, Cell Biology and Metabolism Program, NICHD, Bethesda, MD
Eric Betzig, PhD, Howard Hughes Medical Institute, Janelia Farm Campus, Ashburn, VA
Juan Bonifacino, PhD, Cell Biology and Metabolism Program, NICHD, Bethesda, MD
Michael Davidson, PhD, Florida State University, Tallahassee, FL
Robert DeLotto, PhD, Københavns Universitet, Copenhagen, Denmark
Ellie Ehrenfeld, PhD, Laboratory of Infectious Diseases, NIAID, Bethesda, MD
Ramanuhan Hegde, MD, PhD, Cell Biology and Metabolism Program, NICHD, Bethesda, MD
Harald Hess, PhD, Howard Hughes Medical Institute, Janelia Farm Campus, Ashburn, VA
Cathy Jackson, PhD, Cell Biology and Metabolism Program, NICHD, Bethesda, MD
Mary Lilly, PhD, Cell Biology and Metabolism Program, NICHD, Bethesda, MD
Robert Mullen, PhD, University of Guelph, Guelph, Ontario, Canada
Robert Phair, PhD, BioInformatics, Rockville, MD
Mark Philips, MD, New York University Medical Center, New York, NY
Brian Schaefer, PhD, Uniformed Services University of the Health Sciences, Bethesda, MD
Constantine Stratakis, MD, DSc, Program in Developmental Endocrinology and Genetics, NICHD, Bethesda, MD
Sharon Tooze, PhD, London Research Institute, Cancer Research UK, London, UK
For further information, contact jlippin@helix.nih.gov.
