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Crossing the finish line of development: regulated secretion of Dictyostelium proteins
Crossing the finish line of development: regulated secretion of Dictyostelium proteins
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The entire purpose of the complex programme of multicellular development of the cellular slime mould Dictyostelium discoideum is to convert 80% of the cells into spores that can withstand extremes of temperature and drought. This environmental resistance is due to the thick spore coat, which is a polarized tri-lamellar extracellular matrix made up of glycoproteins on either side of a middle layer of cellulose. Most spore coat glycoproteins are stored in prespore vesicles (PSVs) until an unknown developmental signal triggers them to fuse with the plasma membrane and exocytose their contents for assembly into the spore coats. Regulated protein secretion is important in many developmental events. For example, type II alveolar epithelial cells in the developing lung secrete surfactant proteins1; migrating neural crest cells are guided by many extracellular matrix molecules, including tenascin2 and thrombosphondin-I3, which are secreted by the neural crest cells and surrounding mesenchymal cells of the sclerotome, respectively; and optic nerve growth cone guidance involves an interplay between the extracellular netrin-I and laminin-I proteins that provide positive and negative cues4. However, developmentally regulated protein secretion is rarely as obvious or as amenable to study as in the case of the Dictyostelium prespore cells, where secretion from the PSVs occurs synchronously in 80 000 cells in each multicellular aggregate. Biochemical, molecular genetic, microscopic and proteomic studies are revealing how this pivotal developmental event is controlled.
Supriya Srinivasan, Hannah Alexander and Stephen Alexander The genesis of the spore coat of Dictyostelium represents an exquisite example of developmentally regulated protein secretion. The proteins that are destined to be assembled into the extracellular matrix of the spore coat are stored in unique prespore vesicles that are triggered to secrete their contents at terminal differentiation. The regulation of this process is being revealed by the identification of the individual proteins in these vesicles.
The developmental biology of Dictyostelium discoideum Dictyostelium discoideum can exist indefinitely as free-living amoebae that feed on bacteria or defined medium and divide by mitosis. However, depletion of the food source triggers a dramatic change in the lifestyle of the cells from vegetative growth to multicellular development. There is an abrupt halt in DNA synthesis and cell division, and cells begin to emit pulses of cAMP to which other cells respond by chemotaxis. This results in the formation of multicellular aggregates, each containing 105 cells. Thus, on a 100-mm Petri dish, 109 amoebae will produce approximately 10 000 identical multicellular aggregates that will continue to progress through the remainder of morphogenesis in synchrony (Fig. 1). The structure of these multicellular tissues is maintained by specific cell–cell adhesion molecules and an extracellular matrix that surrounds each aggregate5. Concomitant with the morphological changes, there is a complex programme of gene expression and cytodifferentiation that produces a fruiting body with a multicellular stalk of approximately 20 000 cells, supporting an apical mass of around 80 000 spores (Fig. 1). The spores and stalk cells arise from progenitor cells called prespore and prestalk cells, respectively. The origin of these differentiated cell populations is determined by the position of each particular cell in the cell cycle at the initiation of development, although their fates are not determined irreversibly until later in development6,7. At the slug stage, the cell types are separated spatially, trends in CELL BIOLOGY (Vol. 10) June 2000
with the prespore cells in the posterior and the prestalk cells in the anterior of the slug8. The slug is capable of migrating along gradients of heat and light, and during this time the cells remain as prestalk and prespore cells. When environmental conditions favour culmination (terminal morphogenesis), each cell type embarks on its own specific programme of gene expression, leading to terminal differentiation into the spore and stalk cells9. Each spore consists of a rugged spore coat that encases a single amoeba. This encapsulation results in spores that are resistant to extremes in environmental conditions, allowing the organism to remain dormant for decades, until conditions change to those favouring vegetative growth. At this time, each spore germinates and releases a single amoeba that resumes feeding and cell division. The formation of the spore coat is therefore a crucial event in the life cycle and represents a major evolutionary advantage to the organism. This article aims to highlight recent work on the genesis of this important extracellular matrix and to identify exciting questions for future study. The spore and its coat Spores of haploid strains have a distinctive oval shape and are smaller than the amoebae, although mutants with round spores also exist10. Diploid strains produce spores that are correspondingly
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The authors are in the Division of Biological Sciences, 303 Tucker Hall, University of Missouri, Columbia, MO 65211-7400, USA. E-mail: alexanderst@ missouri.edu
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FIGURE 1 Life cycle of the cellular slime mould Dictyostelium discoideum. Each multicellular aggregate contains 20 000 anterior prestalk cells and 80 000 posterior prespore cells. These differentiate into mature fruiting bodies, each consisting of 80 000 oval spores supported by a stalk made up of 20 000 vacuolated stalk cells. Development – aggregation to fruiting body formation – takes 24 hours and is initiated by starvation.
larger, and the size is diagnostic for ploidy11. Wildtype spores are resistant to 1% Triton X100, and this property can be used to differentiate between spores and cells in a culture. Studies using several different approaches have been conducted to understand the structure and assembly of the spore coat. Biochemical analyses and detailed electron micrographs of the spore coat reveal a tri-lamellar polarized structure made up of approximately equal parts of cellulose and glycoprotein12 (Fig. 2a). The outermost layer of the coat is electron dense and comprises loosely associated proteins that can be removed by treatment with cold sodium dodecyl sulphate. The middle layer is composed of cellulose13,14, and the inner layer contains proteins that are linked covalently and require heat and a denaturing agent for extraction15. A galactose/N-acetyl-galactosamine-containing polysaccharide (GPS) is also part of the spore coat, lying just proximal to the plasma membrane13 (Fig. 2e). Biochemical analysis of spore coats has identified about 10 abundant proteins and several minor spore coat proteins15,16. The posttranslational modifications, including glycosylation and phosphorylation, have been studied extensively for some of these proteins. For a comprehensive list of all the
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PsB complex SpiA Cellulose
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FIGURE 2 (a) Electron micrographs of an oval spore of Dictyostelium discoideum and (b) the tri-lamellar spore coat. Bars, 0.5 mm (a); 0.1 mm (b). The spore coat layers are described in the text and denoted in the figure as follows: 1, outer layer; 2, middle cellulose layer; 3, inner layer; 4, plasma membrane of the dormant amoeba (also see micrographs in Refs 12–14). (c) Phase-contrast and (d) immunofluorescent micrographs of the interspore matrix surrounding a mass of spores in a fruiting body. The interspore matrix is stained with an anti-PsB antibody (MUD102) and detected with a rhodamine-conjugated secondary antibody. Note that individual spores (arrows) do not stain with the anti-PsB antibody because PsB is found on the inner layer of the spore coat. Bar, 5 mm. (e) Schematic presentation of the tri-lamellar spore coat and the underlying plasma membrane of the dormant amoeba.
spore coat proteins, the corresponding antibodies and the associated references see: http://www. biosci.missouri.edu/alexander/spores. Four of the most abundant proteins – SP96, PsB/SP85, SP70 and SP60 – exist in a preassembled multiprotein complex (PsB complex) that is held together by a combination of covalent and noncovalent bonds17,18. Immunodepletion experiments showed that there is additional free SP96 in the cells19. The proteins of the PsB complex are also found in the interspore matrix19 (Fig. 2c, d). The components of the PsB complex trends in CELL BIOLOGY (Vol. 10) June 2000
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are synthesized coordinately in the prespore cells of the slug, and the cognate genes share common promoter elements20. A series of spore coat protein deletion mutants21 was used to define the order of assembly of these proteins into the PsB complex22. In addition, immunostaining of intact and germinated spores demonstrated that SP96 resides only in the outer layer of the coat, whereas PsB/SP85 resides exclusively in the inner layer19. These data imply that the complex is incorporated into the coat with a specific polarity, and probably spans the central cellulose layer. At least one other protein, SpiA, that is not part of the complex, is also localized specifically to the inner layer of the coat23. It is not known how this unique polarity is achieved, and the exact position of each protein within the mature spore coat awaits the generation of specific antibodies to each protein (Fig. 2e). The PsB multiprotein complex has an endogenous cellulose-binding activity, and it was demonstrated that the assembly of the PsB complex was required for the binding activity22. A subsequent report showed that the cysteine-rich C-terminus of recombinant PsB/SP85 protein had cellulose binding activity24. We suggest that the correct positioning of the proteins within the PsB complex allows the exposure of the cellulose-binding site of the PsB/SP85 protein. To our knowledge, this is the first identification of a cellulose binding activity not associated with a cellulase enzyme, and none of the proteins in the complex contains the cellulose-binding domains typical of bacterial cellulases. Studies of mutants with partially assembled PsB complexes that lack cellulose binding activity22 revealed that there is an abnormal assembly of the spore coat, including an increased permeability of the spore coat to fluorescently labelled lectins21, exposure of the PsB/SP85 protein to the outside of the spore and a rapid loss of spore viability25. These data support the idea that the cellulose binding activity of the PsB complex is required for the proper organization of the cellulose, which is essential for the structural organization and integrity of the spore coat and the viability of the spore (Fig. 2e). Although the overall structure of the spore coat is now fairly well understood, important questions remain about the mechanics of its formation: • How are the spore coat proteins and polysaccharides assembled with the correct polarity? • Where does cellulose synthesis take place and how is it coordinated with spore coat assembly? • What enzymes (e.g. disulphide isomerases and other oxidoreductases) are necessary for coat assembly? These questions can be addressed by detailed biophysical and structural studies on the interactions between the spore coat proteins and polysaccharides, and subcellular localization of cellulose synthase and other enzymes necessary for spore coat assembly. PSVs and spore coat proteins Soon after cell aggregation, a specialized secretory organelle – the prespore vesicle (PSV) – appears in the posterior 80% of the cells that are undergoing prespore differentiation26 (Fig. 3a, b). Morphologically, the PSVs trends in CELL BIOLOGY (Vol. 10) June 2000
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FIGURE 3 (a) Phase-contrast and (b) immunofluorescent micrographs of prespore cells from slugs. Multiple prespore vesicles are stained in each permeabilized cell with an antibody to PsB/SP85 [which is in the prespore vesicles (PSVs)] and detected with a rhodamine-conjugated secondary antibody. Bar, 10 mm. (c) Electron micrograph of a purified prespore vesicle showing the electron-dense core of GPS surrounded by an electron-lucent gap, enclosed by a membrane bilayer. Bar, 0.05 mm. (d) Schematic presentation of cellular changes within prespore cells as development proceeds. The PSVs accumulate in prespore cells in slugs. In preculminants, the PSVs move to, and fuse with, the plasma membrane. Finally, during spore formation, the PSV contents are incorporated into the spore coat of the mature spore.
are distinct, consisting of a membrane bilayer surrounding an electron-lucent gap, with a central electron-dense core region made up of GPS27 (Fig. 3c). The origin of the PSVs remains a mystery. It was suggested that they were modified from lysosomes28, but recent analysis of purified PSVs indicates that they lack the lysosomal marker a-mannosidase at all stages of development29. As prespore cell differentiation proceeds, the PSVs increase in size from 150 nm to 450 nm, presumably as the result of the continued accumulation of their contents rather than the fusion of several PSVs, as there is no reduction in the number of PSVs29. This implies the presence of intermediate transport vesicles that remain to be identified. The PsB complex and other spore coat proteins accumulate within the lumen of the PSVs in the prespore cells of the slug18,30. When slugs are induced to culminate, the PSVs move to the plasma membrane, where the two membranes fuse, resulting in exocytosis of the PSV contents (Fig. 3d). By contrast, in slugs that continue to migrate, the PSVs never fuse, supporting the idea that fusion is a signalmediated event26,29. The recent purification of PSVs from different stages of development has led to studies of the secretion
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comment process at the molecular level, which show that, in addition to the PsB multiprotein complex, uncomplexed spore coat proteins, such as SP75, also accumulate in the PSVs19,29. The specific targeting signals used by these proteins for routing to the PSVs are not known. There is no obvious signal sequence shared by any of these proteins, although deletion of the N-terminal half of PsB/SP85 has been shown to preclude localization to the PSVs31. Moreover, it is interesting that not all spore coat proteins are targeted to the PSVs. An important example is a protein component of the inner wall of the spore coat, SpiA, that is synthesized during culmination, but later than the proteins of the PsB complex23, and is not found in the PSVs29. Similarly, the cellulose that is deposited when the spore coat is constructed is also absent from the PSVs12. The gene for the Dictyostelium cellulose synthase has been cloned recently, but the intracellular location of cellulose synthesis is not yet known32. Thus, there appear to be multiple secretory pathways that are used at terminal spore differentiation. Important outstanding questions regarding the PSVs are: • What is the origin of the PSVs during development? • What is the nature of the intermediate post-Golgi vesicles that transport proteins to the PSVs? • What are the molecular targeting signals that direct proteins to the PSVs? • What is the identity of the putative vesicles that mediate SpiA secretion, and is this process regulated or constitutive? Answers to these questions will require the identification of specific membrane proteins of the PSVs, genetic screens for mutants defective in the accumulation of the PsB complex in the PSVs, and the isolation of the intermediate transport vesicles. The coupling of spore differentiation with fruiting body construction The regulated secretion of the PSV contents at the appropriate time in development is mediated by an unknown developmental signal, and is coordinated with fruiting body construction. The conversion of prespore cells to spores occurs as a downward wave as the prespore mass rises up the stalk, and the signal for encapsulation is thought to originate from prestalk cells at the top of the developing culminant33. Upstream components of the developmental signalling pathway that control this process have been identified through genetic studies and are reviewed in detail elsewhere34,35. The first clue as to how cell-type differentiation might be coupled to fruiting body construction came from studies on prespore cells that express a dominant-negative cAMP-dependent protein kinase A (PKA) regulatory subunit, which cannot bind cAMP. The PKA catalytic subunit is always inactive in these cells, and they form fruiting bodies with undifferentiated prespore cells on top of a normal differentiated stalk. Overexpression of the regulatory subunit of PKA arrests development after aggregation, whereas overexpression of its catalytic subunit results in precocious sporulation. Finally, in the presence of
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8-bromo-cAMP, cells of a strain overexpressing the catalytic subunit of PKA can be induced to differentiate into spores in monolayer cultures. The result of this PKA-mediated activity leads to the continuous synthesis of the spore coat proteins that are then routed to the PSVs where they await the signal for secretion. What triggers PSV fusion? The PSVs eventually fuse with the plasma membrane, and therefore the PSVs must have a fusion machinery whose activity is regulated by the culmination signal. Although v- and t-SNAREs are probably involved in the docking of the PSVs at the plasma membrane, additional regulation is necessary for temporal control. A molecule that might fulfil this regulatory role has been identified on the membranes of purified PSVs. PSV membranes contain a small, 21-kDa GTP-binding protein that is first seen at the slug stage of development29, but PSV membranes from early aggregates do not contain this protein. As would be expected of a molecule that regulates fusion in a developmentally controlled manner, its level on the PSV membranes rises as development proceeds, with peak expression at culmination, when the PSVs are just about to fuse with the plasma membrane29. Members of the Rab GTPase family of small GTP-binding proteins are associated with membranes of secretory vesicles, and are known to control the rate and timing of vesicle fusion in yeast and in several mammalian cell types36. Although the identity of the GTP-binding protein associated with the PSVs is not yet known, a Rab GTPase would be an ideal candidate that could transduce the signal to the PSVs in preparation for secretion of the spore coat proteins. Alternatively, the PSV-specific GTPbinding protein might be a Rho or a Rac GTPase, involved in PSV movement along microtubules or the actin cytoskeleton37. Indeed, we have shown that nocodazole, an inhibitor of microtubule formation, also prevents spore differentiation in a dose-dependent manner (S. Srinivasan et al., unpublished). Interestingly, PSVs do not contain clathrin29, although a clathrin-null mutant cannot complete spore differentiation38. In other systems, Annexin VII, a Ca2+-binding protein, is known to regulate vesicle fusion when the intracellular Ca2+ concentration rises39. It is not known whether Annexin VII regulates PSV fusion, but it is important to note that a mutant strain of D. discoideum lacking Annexin VII is defective in sporulation40. Three fundamental questions need to be addressed about the process of developmentally regulated exocytosis from PSVs: • What is the signal that acts directly on the PSVs to trigger secretion? • What molecular transport mechanism do PSVs use to move to the plasma membrane? • What molecules are used for docking and fusion? The future – PSVs and proteomics Many of the questions raised above require further understanding of the protein components of the PSVs. A two-dimensional gel analysis of the purified PSVs showed that they contain approximately trends in CELL BIOLOGY (Vol. 10) June 2000
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80–100 proteins29. Many of these proteins are the structural components of the future spore coat, whereas others might be enzymes required for the actual construction and assembly of the coat. If we are to understand how the developmentally regulated secretion and assembly of PSV contents occurs at a molecular level, these proteins must be identified. The ongoing large-scale proteome project for Dictyostelium41 is making this goal possible. This approach employs two-dimensional gel electrophoresis and peptide mass fingerprinting for identification of individual proteins. As the sequencing of the entire Dictyostelium genome approaches completion, developing a complete protein map of the PSVs is feasible. Functional studies of the individual PSV proteins and their interactions can then be undertaken to enable us to understand how cells cross the finish line of development. References 1 Rooney, S.A. et al. (1994) Molecular and cellular processing of lung surfactant. FASEB J. 8, 957–967 2 Tucker, R.P. and McKay, S.E. (1991) The expression of tenascin by neural crest cells and glia. Development 112, 1031–1039 3 Tucker, R.P. et al. (1999) Thrombospondin-I and neural crest cell migration. Dev. Dyn. 214, 321–322 4 Hopker, V.H. et al. (1999) Growth-cone attraction to netrin-1 is converted to repulsion by laminin-1. Nature 401, 69–73 5 Loomis, W.F. (1982) The Development of Dictyostelium discoideum, Academic Press 6 Gomer, R.H. and Firtel, R.A. (1987) Cell-autonomous determination of cell-type choice in Dictyostelium development by cell cycle phase. Science 237, 758–762 7 Araki, T. et al. (1997) Symmetry breaking in Dictyostelium morphogenesis – evidence that a combination of cell cycle stage and positional information dictates cell fate. Dev. Biol. 192, 645–648 8 Raper, K.B. (1984) The Dictyostelids, Princeton University Press 9 Williams, J. (1995) Morphogenesis in Dictyostelium: new twists to a notso-old tale. Curr. Opin. Genet. Dev. 5, 426–431 10 Williams, K. et al. (1974) Parasexual genetics in Dictyostelium discoideum: mitotic analysis of acriflavin resistance and growth in axenic medium. J. Gen. Microbiol. 84, 59–69 11 Sussman, R. and Sussman, M. (1963) Ploidal inheritance in the slime mould Dictyostelium discoideum: haploidization and genetic segregation of diploid strains. J. Gen. Microbiol. 30, 349–355 12 West, C.M. and Erdos, G.W. (1990) Formation of the Dictyostelium spore coat. Dev. Genet. 11, 492–506 13 Erdos, G.W. and West, C.M. (1989) Formation and organization of the spore coat of Dictyostelium discoideum. Exp. Mycol. 13, 169–182 14 Hemmes, D.E. et al. (1972) Structural and enzymatic analysis of the spore wall layers in Dictyostelium discoideum. J. Ultrastruct. Res. 41, 406–417 15 Wilkinson, D. et al. (1983) Synthesis of spore proteins during development of Dictyostelium discoideum. Biochem. J. 216, 567–574 16 Orlowski, M. and Loomis, W.F. (1979) Plasma membrane proteins of Dictyostelium: the spore coat proteins. Dev. Biol. 71, 297–307 17 Devine, K.M. et al. (1982) Differential synthesis of spore coat proteins in prespore and prestalk cells of Dictyostelium. Proc. Natl. Acad. Sci. U. S. A. 79, 7361–7365
18 Watson, N. et al. (1993) A developmentally regulated glycoprotein complex from Dictyostelium discoideum. J. Biol. Chem. 268, 22634–22641 19 Watson, N. et al. (1994) The PsB complex is secreted as a preassembled precursor of the spore coat in Dictyostelium discoideum. J. Cell Sci. 107, 2567–2579 20 Fosnaugh, K.L. and Loomis, W.F. (1991) Coordinate regulation of the spore coat genes in Dictyostelium discoideum. Dev. Genet. 12, 123–132 21 Fosnaugh, K.L. et al. (1994) Structural roles of the spore coat proteins in Dictyostelium discoideum. Dev. Biol. 166, 823–825 22 McGuire, V. and Alexander, S. (1996) PsB multiprotein complex of Dictyostelium discoideum: demonstration of cellulose binding activity and order of protein subunit assembly. J. Biol. Chem. 271, 14596–14603 23 Richardson, D.L. and Loomis, W.F. (1992) Disruption of the sporulationspecific gene spiA in Dictyostelium discoideum leads to spore instability. Genes Dev. 6, 1058–1070 24 Zhang, Y.Y. et al. (1998) Two proteins of the Dictyostelium spore coat bind to cellulose in vitro. Biochemistry 37, 10766–10779 25 Srinivasan, S. et al. The cellulose binding activity of the PsB multiprotein complex is required for proper assembly of the spore coat and spore viability in Dictyostelium discoideum (in press) 26 Hohl, H. and Hamamoto, S. (1969) Ultrastructure of spore differentiation in Dictyostelium: the prespore vacuole. J. Ultrastruct. Res. 26, 442–453 27 Maeda, Y. and Takeuchi, I. (1969) Cell differentiation and fine structures in the development of the cellular slime molds. Dev. Growth Differ. 11, 232–245 28 Lenhard, J.M. et al. (1990) Developing Dictyostelium cells contain the lysosomal enzyme alpha-mannosidase in a secretory granule. J. Cell Biol. 109, 2761–2769 29 Srinivasan, S. et al. (1999) The prespore vesicles of Dictyostelium discoideum: purification, characterization and developmental regulation. J. Biol. Chem. 274, 35823–35831 30 Devine, K.M. et al. (1983) Spore coat proteins of Dictyostelium discoideum are packaged in prespore vesicles. Dev. Biol. 99, 437–446 31 Zhang, Y. et al. (1999) A linking function for the cellulose-binding protein SP85 in the spore coat of Dictyostelium discoideum. J. Cell Sci. 112, 4367–4377 32 Blanton, R.L. et al. (2000) The cellulose synthase gene of Dictyostelium. Proc. Natl. Acad. Sci. U. S. A. 97, 2391–2396 33 Loomis, W.F. et al. (1998) Two-component signal transduction systems in eukaryotic microorganisms. Curr. Opin. Microbiol. 1, 643–648 34 Loomis, W.F. (1998) Role of PKA in the timing of developmental events in Dictyostelium cells. Microbiol. Mol. Biol. Rev. 62, 684–694 35 Thomason, P. et al. (1999) Taking the plunge – terminal differentiation in Dictyostelium. Trends Genet. 15, 15–19 36 Novick, P. and Zerial, M. (1997) The diversity of Rab proteins in vesicle transport. Curr. Opin. Cell Biol. 9, 496–504 37 Clague, M.J. (1998) Molecular aspects of the endocytic pathway. Biochem. J. 336, 271–282 38 Niswonger, M.L. and O’Halloran, T.J. (1997) Clathrin heavy chain is required for spore cell but not stalk cell differentiation in Dictyostelium discoideum. Development 124, 443–451 39 Donnelly, S.R. and Moss, S.E. (1997) Annexins in the secretory pathway. Cell. Mol. Life Sci. 53, 533–538 40 Döring, V. et al. (1995) The in vivo role of annexin VII (synexin): characterization of an annexin VII-deficient Dictyostelium mutant indicates an involvement in Ca2+-regulated processes. J. Cell Sci. 108, 2065–2076 41 Yan, J.X. et al. (1997) The Dictyostelium proteome project: studying the readout of the genome. In Dictyostelium: a Model System for Cell and Developmental Biology (Maeda, Y. et al., eds), pp. 455–469, Universal Academy Press
Acknowledgements The work from this laboratory was supported by a University of Missouri Research Board Grant RB97-044. S.S. received fellowship support from the Genetics Area Program of the University of Missouri, and this work is in partial fulfilment of the requirements for a PhD in this program. S.A. is the recipient of an American Cancer Society Faculty Research Award FRA-448. We thank Professors Steve Nothwehr, Keith Williams and Cathy Krull for discussions and comments on the manuscript, and M. Xenia U. Garcia, Guochun Li and Christopher Foote for helpful discussions.
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trends in CELL BIOLOGY (Vol. 10) June 2000
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The entire purpose of the complex programme of multicellular development of the cellular slime mould Dictyostelium discoideum is to convert 80% of the cells into spores that can withstand extremes of temperature and drought. This environmental resistance is due to the thick spore coat, which is a polarized tri-lamellar extracellular matrix made up of glycoproteins on either side of a middle layer of cellulose. Most spore coat glycoproteins are stored in prespore vesicles (PSVs) until an unknown developmental signal triggers them to fuse with the plasma membrane and exocytose their contents for assembly into the spore coats. Regulated protein secretion is important in many developmental events. For example, type II alveolar epithelial cells in the developing lung secrete surfactant proteins1; migrating neural crest cells are guided by many extracellular matrix molecules, including tenascin2 and thrombosphondin-I3, which are secreted by the neural crest cells and surrounding mesenchymal cells of the sclerotome, respectively; and optic nerve growth cone guidance involves an interplay between the extracellular netrin-I and laminin-I proteins that provide positive and negative cues4. However, developmentally regulated protein secretion is rarely as obvious or as amenable to study as in the case of the Dictyostelium prespore cells, where secretion from the PSVs occurs synchronously in 80 000 cells in each multicellular aggregate. Biochemical, molecular genetic, microscopic and proteomic studies are revealing how this pivotal developmental event is controlled.
Supriya Srinivasan, Hannah Alexander and Stephen Alexander The genesis of the spore coat of Dictyostelium represents an exquisite example of developmentally regulated protein secretion. The proteins that are destined to be assembled into the extracellular matrix of the spore coat are stored in unique prespore vesicles that are triggered to secrete their contents at terminal differentiation. The regulation of this process is being revealed by the identification of the individual proteins in these vesicles.
The developmental biology of Dictyostelium discoideum Dictyostelium discoideum can exist indefinitely as free-living amoebae that feed on bacteria or defined medium and divide by mitosis. However, depletion of the food source triggers a dramatic change in the lifestyle of the cells from vegetative growth to multicellular development. There is an abrupt halt in DNA synthesis and cell division, and cells begin to emit pulses of cAMP to which other cells respond by chemotaxis. This results in the formation of multicellular aggregates, each containing 105 cells. Thus, on a 100-mm Petri dish, 109 amoebae will produce approximately 10 000 identical multicellular aggregates that will continue to progress through the remainder of morphogenesis in synchrony (Fig. 1). The structure of these multicellular tissues is maintained by specific cell–cell adhesion molecules and an extracellular matrix that surrounds each aggregate5. Concomitant with the morphological changes, there is a complex programme of gene expression and cytodifferentiation that produces a fruiting body with a multicellular stalk of approximately 20 000 cells, supporting an apical mass of around 80 000 spores (Fig. 1). The spores and stalk cells arise from progenitor cells called prespore and prestalk cells, respectively. The origin of these differentiated cell populations is determined by the position of each particular cell in the cell cycle at the initiation of development, although their fates are not determined irreversibly until later in development6,7. At the slug stage, the cell types are separated spatially, trends in CELL BIOLOGY (Vol. 10) June 2000
with the prespore cells in the posterior and the prestalk cells in the anterior of the slug8. The slug is capable of migrating along gradients of heat and light, and during this time the cells remain as prestalk and prespore cells. When environmental conditions favour culmination (terminal morphogenesis), each cell type embarks on its own specific programme of gene expression, leading to terminal differentiation into the spore and stalk cells9. Each spore consists of a rugged spore coat that encases a single amoeba. This encapsulation results in spores that are resistant to extremes in environmental conditions, allowing the organism to remain dormant for decades, until conditions change to those favouring vegetative growth. At this time, each spore germinates and releases a single amoeba that resumes feeding and cell division. The formation of the spore coat is therefore a crucial event in the life cycle and represents a major evolutionary advantage to the organism. This article aims to highlight recent work on the genesis of this important extracellular matrix and to identify exciting questions for future study. The spore and its coat Spores of haploid strains have a distinctive oval shape and are smaller than the amoebae, although mutants with round spores also exist10. Diploid strains produce spores that are correspondingly
0962-8924/00/$ – see front matter © 2000 Elsevier Science Ltd. All rights reserved. PII: S962-8924(00)01758-X
The authors are in the Division of Biological Sciences, 303 Tucker Hall, University of Missouri, Columbia, MO 65211-7400, USA. E-mail: alexanderst@ missouri.edu
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trends in Cell Biology
FIGURE 1 Life cycle of the cellular slime mould Dictyostelium discoideum. Each multicellular aggregate contains 20 000 anterior prestalk cells and 80 000 posterior prespore cells. These differentiate into mature fruiting bodies, each consisting of 80 000 oval spores supported by a stalk made up of 20 000 vacuolated stalk cells. Development – aggregation to fruiting body formation – takes 24 hours and is initiated by starvation.
larger, and the size is diagnostic for ploidy11. Wildtype spores are resistant to 1% Triton X100, and this property can be used to differentiate between spores and cells in a culture. Studies using several different approaches have been conducted to understand the structure and assembly of the spore coat. Biochemical analyses and detailed electron micrographs of the spore coat reveal a tri-lamellar polarized structure made up of approximately equal parts of cellulose and glycoprotein12 (Fig. 2a). The outermost layer of the coat is electron dense and comprises loosely associated proteins that can be removed by treatment with cold sodium dodecyl sulphate. The middle layer is composed of cellulose13,14, and the inner layer contains proteins that are linked covalently and require heat and a denaturing agent for extraction15. A galactose/N-acetyl-galactosamine-containing polysaccharide (GPS) is also part of the spore coat, lying just proximal to the plasma membrane13 (Fig. 2e). Biochemical analysis of spore coats has identified about 10 abundant proteins and several minor spore coat proteins15,16. The posttranslational modifications, including glycosylation and phosphorylation, have been studied extensively for some of these proteins. For a comprehensive list of all the
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PsB complex SpiA Cellulose
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FIGURE 2 (a) Electron micrographs of an oval spore of Dictyostelium discoideum and (b) the tri-lamellar spore coat. Bars, 0.5 mm (a); 0.1 mm (b). The spore coat layers are described in the text and denoted in the figure as follows: 1, outer layer; 2, middle cellulose layer; 3, inner layer; 4, plasma membrane of the dormant amoeba (also see micrographs in Refs 12–14). (c) Phase-contrast and (d) immunofluorescent micrographs of the interspore matrix surrounding a mass of spores in a fruiting body. The interspore matrix is stained with an anti-PsB antibody (MUD102) and detected with a rhodamine-conjugated secondary antibody. Note that individual spores (arrows) do not stain with the anti-PsB antibody because PsB is found on the inner layer of the spore coat. Bar, 5 mm. (e) Schematic presentation of the tri-lamellar spore coat and the underlying plasma membrane of the dormant amoeba.
spore coat proteins, the corresponding antibodies and the associated references see: http://www. biosci.missouri.edu/alexander/spores. Four of the most abundant proteins – SP96, PsB/SP85, SP70 and SP60 – exist in a preassembled multiprotein complex (PsB complex) that is held together by a combination of covalent and noncovalent bonds17,18. Immunodepletion experiments showed that there is additional free SP96 in the cells19. The proteins of the PsB complex are also found in the interspore matrix19 (Fig. 2c, d). The components of the PsB complex trends in CELL BIOLOGY (Vol. 10) June 2000
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are synthesized coordinately in the prespore cells of the slug, and the cognate genes share common promoter elements20. A series of spore coat protein deletion mutants21 was used to define the order of assembly of these proteins into the PsB complex22. In addition, immunostaining of intact and germinated spores demonstrated that SP96 resides only in the outer layer of the coat, whereas PsB/SP85 resides exclusively in the inner layer19. These data imply that the complex is incorporated into the coat with a specific polarity, and probably spans the central cellulose layer. At least one other protein, SpiA, that is not part of the complex, is also localized specifically to the inner layer of the coat23. It is not known how this unique polarity is achieved, and the exact position of each protein within the mature spore coat awaits the generation of specific antibodies to each protein (Fig. 2e). The PsB multiprotein complex has an endogenous cellulose-binding activity, and it was demonstrated that the assembly of the PsB complex was required for the binding activity22. A subsequent report showed that the cysteine-rich C-terminus of recombinant PsB/SP85 protein had cellulose binding activity24. We suggest that the correct positioning of the proteins within the PsB complex allows the exposure of the cellulose-binding site of the PsB/SP85 protein. To our knowledge, this is the first identification of a cellulose binding activity not associated with a cellulase enzyme, and none of the proteins in the complex contains the cellulose-binding domains typical of bacterial cellulases. Studies of mutants with partially assembled PsB complexes that lack cellulose binding activity22 revealed that there is an abnormal assembly of the spore coat, including an increased permeability of the spore coat to fluorescently labelled lectins21, exposure of the PsB/SP85 protein to the outside of the spore and a rapid loss of spore viability25. These data support the idea that the cellulose binding activity of the PsB complex is required for the proper organization of the cellulose, which is essential for the structural organization and integrity of the spore coat and the viability of the spore (Fig. 2e). Although the overall structure of the spore coat is now fairly well understood, important questions remain about the mechanics of its formation: • How are the spore coat proteins and polysaccharides assembled with the correct polarity? • Where does cellulose synthesis take place and how is it coordinated with spore coat assembly? • What enzymes (e.g. disulphide isomerases and other oxidoreductases) are necessary for coat assembly? These questions can be addressed by detailed biophysical and structural studies on the interactions between the spore coat proteins and polysaccharides, and subcellular localization of cellulose synthase and other enzymes necessary for spore coat assembly. PSVs and spore coat proteins Soon after cell aggregation, a specialized secretory organelle – the prespore vesicle (PSV) – appears in the posterior 80% of the cells that are undergoing prespore differentiation26 (Fig. 3a, b). Morphologically, the PSVs trends in CELL BIOLOGY (Vol. 10) June 2000
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FIGURE 3 (a) Phase-contrast and (b) immunofluorescent micrographs of prespore cells from slugs. Multiple prespore vesicles are stained in each permeabilized cell with an antibody to PsB/SP85 [which is in the prespore vesicles (PSVs)] and detected with a rhodamine-conjugated secondary antibody. Bar, 10 mm. (c) Electron micrograph of a purified prespore vesicle showing the electron-dense core of GPS surrounded by an electron-lucent gap, enclosed by a membrane bilayer. Bar, 0.05 mm. (d) Schematic presentation of cellular changes within prespore cells as development proceeds. The PSVs accumulate in prespore cells in slugs. In preculminants, the PSVs move to, and fuse with, the plasma membrane. Finally, during spore formation, the PSV contents are incorporated into the spore coat of the mature spore.
are distinct, consisting of a membrane bilayer surrounding an electron-lucent gap, with a central electron-dense core region made up of GPS27 (Fig. 3c). The origin of the PSVs remains a mystery. It was suggested that they were modified from lysosomes28, but recent analysis of purified PSVs indicates that they lack the lysosomal marker a-mannosidase at all stages of development29. As prespore cell differentiation proceeds, the PSVs increase in size from 150 nm to 450 nm, presumably as the result of the continued accumulation of their contents rather than the fusion of several PSVs, as there is no reduction in the number of PSVs29. This implies the presence of intermediate transport vesicles that remain to be identified. The PsB complex and other spore coat proteins accumulate within the lumen of the PSVs in the prespore cells of the slug18,30. When slugs are induced to culminate, the PSVs move to the plasma membrane, where the two membranes fuse, resulting in exocytosis of the PSV contents (Fig. 3d). By contrast, in slugs that continue to migrate, the PSVs never fuse, supporting the idea that fusion is a signalmediated event26,29. The recent purification of PSVs from different stages of development has led to studies of the secretion
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8-bromo-cAMP, cells of a strain overexpressing the catalytic subunit of PKA can be induced to differentiate into spores in monolayer cultures. The result of this PKA-mediated activity leads to the continuous synthesis of the spore coat proteins that are then routed to the PSVs where they await the signal for secretion. What triggers PSV fusion? The PSVs eventually fuse with the plasma membrane, and therefore the PSVs must have a fusion machinery whose activity is regulated by the culmination signal. Although v- and t-SNAREs are probably involved in the docking of the PSVs at the plasma membrane, additional regulation is necessary for temporal control. A molecule that might fulfil this regulatory role has been identified on the membranes of purified PSVs. PSV membranes contain a small, 21-kDa GTP-binding protein that is first seen at the slug stage of development29, but PSV membranes from early aggregates do not contain this protein. As would be expected of a molecule that regulates fusion in a developmentally controlled manner, its level on the PSV membranes rises as development proceeds, with peak expression at culmination, when the PSVs are just about to fuse with the plasma membrane29. Members of the Rab GTPase family of small GTP-binding proteins are associated with membranes of secretory vesicles, and are known to control the rate and timing of vesicle fusion in yeast and in several mammalian cell types36. Although the identity of the GTP-binding protein associated with the PSVs is not yet known, a Rab GTPase would be an ideal candidate that could transduce the signal to the PSVs in preparation for secretion of the spore coat proteins. Alternatively, the PSV-specific GTPbinding protein might be a Rho or a Rac GTPase, involved in PSV movement along microtubules or the actin cytoskeleton37. Indeed, we have shown that nocodazole, an inhibitor of microtubule formation, also prevents spore differentiation in a dose-dependent manner (S. Srinivasan et al., unpublished). Interestingly, PSVs do not contain clathrin29, although a clathrin-null mutant cannot complete spore differentiation38. In other systems, Annexin VII, a Ca2+-binding protein, is known to regulate vesicle fusion when the intracellular Ca2+ concentration rises39. It is not known whether Annexin VII regulates PSV fusion, but it is important to note that a mutant strain of D. discoideum lacking Annexin VII is defective in sporulation40. Three fundamental questions need to be addressed about the process of developmentally regulated exocytosis from PSVs: • What is the signal that acts directly on the PSVs to trigger secretion? • What molecular transport mechanism do PSVs use to move to the plasma membrane? • What molecules are used for docking and fusion? The future – PSVs and proteomics Many of the questions raised above require further understanding of the protein components of the PSVs. A two-dimensional gel analysis of the purified PSVs showed that they contain approximately trends in CELL BIOLOGY (Vol. 10) June 2000
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80–100 proteins29. Many of these proteins are the structural components of the future spore coat, whereas others might be enzymes required for the actual construction and assembly of the coat. If we are to understand how the developmentally regulated secretion and assembly of PSV contents occurs at a molecular level, these proteins must be identified. The ongoing large-scale proteome project for Dictyostelium41 is making this goal possible. This approach employs two-dimensional gel electrophoresis and peptide mass fingerprinting for identification of individual proteins. As the sequencing of the entire Dictyostelium genome approaches completion, developing a complete protein map of the PSVs is feasible. Functional studies of the individual PSV proteins and their interactions can then be undertaken to enable us to understand how cells cross the finish line of development. References 1 Rooney, S.A. et al. (1994) Molecular and cellular processing of lung surfactant. FASEB J. 8, 957–967 2 Tucker, R.P. and McKay, S.E. (1991) The expression of tenascin by neural crest cells and glia. Development 112, 1031–1039 3 Tucker, R.P. et al. (1999) Thrombospondin-I and neural crest cell migration. Dev. Dyn. 214, 321–322 4 Hopker, V.H. et al. (1999) Growth-cone attraction to netrin-1 is converted to repulsion by laminin-1. Nature 401, 69–73 5 Loomis, W.F. (1982) The Development of Dictyostelium discoideum, Academic Press 6 Gomer, R.H. and Firtel, R.A. (1987) Cell-autonomous determination of cell-type choice in Dictyostelium development by cell cycle phase. Science 237, 758–762 7 Araki, T. et al. (1997) Symmetry breaking in Dictyostelium morphogenesis – evidence that a combination of cell cycle stage and positional information dictates cell fate. Dev. Biol. 192, 645–648 8 Raper, K.B. (1984) The Dictyostelids, Princeton University Press 9 Williams, J. (1995) Morphogenesis in Dictyostelium: new twists to a notso-old tale. Curr. Opin. Genet. Dev. 5, 426–431 10 Williams, K. et al. (1974) Parasexual genetics in Dictyostelium discoideum: mitotic analysis of acriflavin resistance and growth in axenic medium. J. Gen. Microbiol. 84, 59–69 11 Sussman, R. and Sussman, M. (1963) Ploidal inheritance in the slime mould Dictyostelium discoideum: haploidization and genetic segregation of diploid strains. J. Gen. Microbiol. 30, 349–355 12 West, C.M. and Erdos, G.W. (1990) Formation of the Dictyostelium spore coat. Dev. Genet. 11, 492–506 13 Erdos, G.W. and West, C.M. (1989) Formation and organization of the spore coat of Dictyostelium discoideum. Exp. Mycol. 13, 169–182 14 Hemmes, D.E. et al. (1972) Structural and enzymatic analysis of the spore wall layers in Dictyostelium discoideum. J. Ultrastruct. Res. 41, 406–417 15 Wilkinson, D. et al. (1983) Synthesis of spore proteins during development of Dictyostelium discoideum. Biochem. J. 216, 567–574 16 Orlowski, M. and Loomis, W.F. (1979) Plasma membrane proteins of Dictyostelium: the spore coat proteins. Dev. Biol. 71, 297–307 17 Devine, K.M. et al. (1982) Differential synthesis of spore coat proteins in prespore and prestalk cells of Dictyostelium. Proc. Natl. Acad. Sci. U. S. A. 79, 7361–7365
18 Watson, N. et al. (1993) A developmentally regulated glycoprotein complex from Dictyostelium discoideum. J. Biol. Chem. 268, 22634–22641 19 Watson, N. et al. (1994) The PsB complex is secreted as a preassembled precursor of the spore coat in Dictyostelium discoideum. J. Cell Sci. 107, 2567–2579 20 Fosnaugh, K.L. and Loomis, W.F. (1991) Coordinate regulation of the spore coat genes in Dictyostelium discoideum. Dev. Genet. 12, 123–132 21 Fosnaugh, K.L. et al. (1994) Structural roles of the spore coat proteins in Dictyostelium discoideum. Dev. Biol. 166, 823–825 22 McGuire, V. and Alexander, S. (1996) PsB multiprotein complex of Dictyostelium discoideum: demonstration of cellulose binding activity and order of protein subunit assembly. J. Biol. Chem. 271, 14596–14603 23 Richardson, D.L. and Loomis, W.F. (1992) Disruption of the sporulationspecific gene spiA in Dictyostelium discoideum leads to spore instability. Genes Dev. 6, 1058–1070 24 Zhang, Y.Y. et al. (1998) Two proteins of the Dictyostelium spore coat bind to cellulose in vitro. Biochemistry 37, 10766–10779 25 Srinivasan, S. et al. The cellulose binding activity of the PsB multiprotein complex is required for proper assembly of the spore coat and spore viability in Dictyostelium discoideum (in press) 26 Hohl, H. and Hamamoto, S. (1969) Ultrastructure of spore differentiation in Dictyostelium: the prespore vacuole. J. Ultrastruct. Res. 26, 442–453 27 Maeda, Y. and Takeuchi, I. (1969) Cell differentiation and fine structures in the development of the cellular slime molds. Dev. Growth Differ. 11, 232–245 28 Lenhard, J.M. et al. (1990) Developing Dictyostelium cells contain the lysosomal enzyme alpha-mannosidase in a secretory granule. J. Cell Biol. 109, 2761–2769 29 Srinivasan, S. et al. (1999) The prespore vesicles of Dictyostelium discoideum: purification, characterization and developmental regulation. J. Biol. Chem. 274, 35823–35831 30 Devine, K.M. et al. (1983) Spore coat proteins of Dictyostelium discoideum are packaged in prespore vesicles. Dev. Biol. 99, 437–446 31 Zhang, Y. et al. (1999) A linking function for the cellulose-binding protein SP85 in the spore coat of Dictyostelium discoideum. J. Cell Sci. 112, 4367–4377 32 Blanton, R.L. et al. (2000) The cellulose synthase gene of Dictyostelium. Proc. Natl. Acad. Sci. U. S. A. 97, 2391–2396 33 Loomis, W.F. et al. (1998) Two-component signal transduction systems in eukaryotic microorganisms. Curr. Opin. Microbiol. 1, 643–648 34 Loomis, W.F. (1998) Role of PKA in the timing of developmental events in Dictyostelium cells. Microbiol. Mol. Biol. Rev. 62, 684–694 35 Thomason, P. et al. (1999) Taking the plunge – terminal differentiation in Dictyostelium. Trends Genet. 15, 15–19 36 Novick, P. and Zerial, M. (1997) The diversity of Rab proteins in vesicle transport. Curr. Opin. Cell Biol. 9, 496–504 37 Clague, M.J. (1998) Molecular aspects of the endocytic pathway. Biochem. J. 336, 271–282 38 Niswonger, M.L. and O’Halloran, T.J. (1997) Clathrin heavy chain is required for spore cell but not stalk cell differentiation in Dictyostelium discoideum. Development 124, 443–451 39 Donnelly, S.R. and Moss, S.E. (1997) Annexins in the secretory pathway. Cell. Mol. Life Sci. 53, 533–538 40 Döring, V. et al. (1995) The in vivo role of annexin VII (synexin): characterization of an annexin VII-deficient Dictyostelium mutant indicates an involvement in Ca2+-regulated processes. J. Cell Sci. 108, 2065–2076 41 Yan, J.X. et al. (1997) The Dictyostelium proteome project: studying the readout of the genome. In Dictyostelium: a Model System for Cell and Developmental Biology (Maeda, Y. et al., eds), pp. 455–469, Universal Academy Press
Acknowledgements The work from this laboratory was supported by a University of Missouri Research Board Grant RB97-044. S.S. received fellowship support from the Genetics Area Program of the University of Missouri, and this work is in partial fulfilment of the requirements for a PhD in this program. S.A. is the recipient of an American Cancer Society Faculty Research Award FRA-448. We thank Professors Steve Nothwehr, Keith Williams and Cathy Krull for discussions and comments on the manuscript, and M. Xenia U. Garcia, Guochun Li and Christopher Foote for helpful discussions.
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