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The identification of Dictyostelium phosphoproteins altered in response to the activation of RasG Secko, David Matthew 2004

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THE IDENTIFICATION OF DICTYOSTELIUM PHOSPHOPROTEINS ALTERED IN RESPONSE TO THE ACTIVATION OF RASG By DAVID MATTHEW SECKO B.Sc. (Hons), Queen's University, 1998 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY In THE FACULTY OF GRADUATE STUDIES (Department of Microbiology and Immunology) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA June 2004 © David Matthew Secko, 2004 THE IDENTIFICATION OF DICTYOSTELIUM PHOSPHOPROTIENS ALTERED IN RESPONSE TO THE ACTIVATION OF RASG By DAVID MATTHEW SECKO B.Sc. (Hons), Queen's University, 1998 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY In T H E FACULTY OF GRADUATE STUDIES (Department of Microbiology and Immunology) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA June 2004 © David Matthew Secko, 2004 A B S T R A C T Dictyosteliiim RasG has been implicated in the regulation of a variety of cellular processes, including the initiation of development, cell movement, and cytokinesis, but the molecular components of the signaling pathways involved are largely unknown. This thesis describes the search for putative downstream targets of the RasG signaling pathway through the identification of proteins whose level of phosphorylation were changed in response to the induction of a gene encoding an activated form of RasG, RasG(G12T). Two expression systems were tested to increase the level of RasG(G12T) in the cell. One system used the rasG(G12T) gene fused to the ribonucleotide reductase (rnrB) promoter. Induction of RasG(G12T) expression from this promoter was accompanied by a decrease in discoidin mRNA levels, a proposed negatively regulated target of RasG. Regulation of Discoidin expression was also down regulated in rasG null cells under all experimental conditions, but the response to well established discoidin regulators still occurred in this strain. These results revealed a role for RasG in modulating discoidin gene expression. A tetracycline-regulated expression system was used to study the effect of increasing the level of RasG(G12T) on the phosphorylation state of Dictyostelium proteins. Over 70 phosphorylated proteins in vegetative cells were resolved by 2D immunoblot analysis and thirteen proteins, which reproducibly changed in response to RasG(G12T) expression, were recovered from 2D gels and identified by mass spectrometry. The proteins identified were: the signaling proteins RasGEF-R and protein kinase B (PKB), the adhesion protein DdCAD-1, the cytoskeletal protein actin, the mitochondrial division protein FtsZA, as well as several proteins involved in protein translation or metabolism. An additional set of experiments showed that phosphorlyation of the vacuolar H+-ATPase component VatA upon folate stimulation was modulated by RasG. Two of the proteins whose phosphorylation levels were affected by RasG(G12T), DdCAD-1 and PKB, were analyzed further. Cells expressing RasG(G12T) exhibited increased cohesion and this increase correlated with DdCAD-1 localization and dephosphorylation. Induction of RasG(G12T) was also found to up-regulate both the basal and folate-induced level of phosphorylation of PKB, and cells expressing RasG(G12T) contained slightly more membrane bound P K B . Together the presented work has identified several proteins whose phosphorylation state was affected by RasG(G12T) and additional regulatory roles for RasG. 11 T A B L E O F CONTENTS A B S T R A C T i i T A B L E O F CONTENTS i i i LIST O F T A B L E S vii LIST O F FIGURES viii LIST O F ABBREVIATIONS x A C K N O W L E D G E M E N T S xii 1 INTRODUCTION 1 1.1 A historical introduction to Ras research 1 1.2 An overview of Ras biology 1 1.2.1 The Ras superfamily of proteins 2 1.2.2 The GTP/GDP switch 3 1.2.3 Ras activation 3 1.2.4 Ras effectors 5 1.3 A broader understanding of Ras function through model organisms 9 1.4 Dictyostelium as a model organism 9 1.5 Dictyostelium Ras proteins 13 1.6 Thesis objectives 16 1.6.1 Background 16 1.6.2 Objectives 17 2 M A T E R I A L S AND M E T H O D S 18 2.1 Materials 18 2.1.1 Reagents 18 2.1.2 Services 18 2.1.3 Dictyostelium strains 19 2.1.4 Plasmids 19 2.2 Methods 19 2.2.1 Dictyostelium cell culture 19 2.2.2 Dictyostelium growth in bacterial suspension..... 20 2.2.3 Dictyostelium development 20 ii i 2.2.4 Vector constructions and Polymerase Chain Reaction (PCR) 20 2.2.5 Dictyostelium transformations 22 2.2.6 Induction conditions for protein overexpression 23 2.2.7 R N A analysis 23 2.2.8 One dimensional (ID) immunoblot analysis 24 2.2.9 Two dimensional (2D) electrophoresis and immunoblot analysis 25 2.2.10 In-gel tryptic protein digestion 27 2.2.11 Mass spectral identification and analysis 28 2.2.12 Microscopy 28 2.2.13 Indirect immunofluorescence 28 2.2.14 Cell cohesion assays 30 2.2.15 Chemotaxis and aggregation streaming assays 31 2.2.16 Assay of folate stimulation of ERK2 and P K B phosphorylation 31 2.2.17 Protein translocation assays 31 2.2.18 Subcellular fractionation 32 3 RESULTS , 33 3.1 The effects of RasG proteins levels on discoidin expression 33 3.1.1 Introduction 33 3.1.2 Accumulation of RasG(G12T) is accompanied by decreased discoidin mRNA levels 33 3.1.3 Discoidin expression in rasG null cells 33 3.1.4 Summary 38 3.2 Use of an inducible expression system to identify Dictyostelium phosphoproteins altered by the expression of RasG(G12T) 41 3.2.1 Introduction 41 3.2.2 Expression of RasG(G12T) from the tetracycline promoter, an alternative expression system 41 3.2.3 Detection of protein phosphorylation changes upon the expression of activated RasG 43 3.2.4 Separation of RasG-responsive phosphoproteins by 2D gel electrophoresis 43 iv 3.2.5 Identification of putative RasG-responsive phosphoproteins 49 3.2.6 Total protein level of a subset of RasG-responsive phosphoproteins does not change upon RasG(G12T) expression 52 3.2.7 Confirmation of the identities of RasGEF-R and DdCAD-1 53 3.2.8 Summary 57 3.3 Activated RasG and cell-to-cell adhesion in Dictyostelium 58 3.3.1 Introduction 58 3.3.2 Expression of RasG(G 12T) during vegetative growth is accompanied by cell clumping 58 3.3.3 RasG(G12T)-induced vegetative cell clumping can be blocked with DdCAD-1 inhibitors 58 3.3.4 DdCAD-1 localizes to the cell surface upon the expression of RasG(G12T) 60 3.3.5 Cell adhesion during early developmental is enhanced in RasG(G12T) expressing cells 63 3.3.6 DdCAD-1 phosphorylation decreases during Dictyostelium development 63 3.3.7 A correlation between the decrease in DdCAD-1 phosphorylation and the cellular localization of DdCAD-1 65 3.3.8 rasG null cells are defective in streaming during aggregation 68 3.3.9 RasG is required for high levels of DdCAD-1 expression 71 3.3.10 Summary 71 3 .4 Effect of R a s G ( G 1 2 T ) on Dictyostelium PKB phosphorylation 73 3.4.1 Introduction 73 3.4.2 Enhancement of folate-induced phosphorylation of Dictyostelium P K B in cells expressing RasG(G12T) 73 3.4.3 Cells expressing RasG(Gl 2T) have more P K B in their membrane fractions 76 3.4.4 Cell polarity in RasG(G 12T) expressing cells 77 3.4.5 Summary 80 3.5 Identification of a protein phosphorylation in response to a chemoattractant and its modulation by RasG 81 3.5.1 Introduction , 81 3.5.2 Use of narrow-range 2D gel electrophoresis for the resolution of p70 81 3.5.3 LrrA is a predicted Ras-binding protein, but is not p70 84 3.5.4 Isolation of a protein from the IrrA null that co-migrates with LrrA in wild type cells 90 3.5.5 Attempts at confirming that p70 is VatA 90 3.5.6 Summary 95 4 DISCUSSION 96 4.1 Inducible expression of activated RasG and its effect on discoidin expression 96 4.2 Combining RasG activation and functional proteomics to uncover novel phosphoproteins 98 4.3 The role of Dictyostelium kinases and phosphatases 101 4.4 Actin phosphorylation and RasG regulation of the cytoskeleton 104 4.5 Which RasGEF is the right RasGEF for RasG? 105 4.6 A role for RasG in regulating early developmental adhesion through the dephosphorylation of DdCAD-1 107 4.7 Effect of RasG(G12T) on P K B phosphorylation 110 4.8 Additional targets of RasG regulation 113 4.9 Emerging themes of the RasG signaling pathway(s) and future prospects .... 114 BIBLIOGRAPHY 116 LIST O F T A B L E S Table 1: A list of Ras-responsive phosphoproteins identified in this study 51 Table 2: A list of proteins identified as putative p70 candidates 84 vii LIST O F FIGURES Figure 1. A model of the Ras GTP/GDP switch 4 Figure 2. Downstream effectors of Ras proteins and their functions 7 Figure 3. Multicellular development of Dictyostelium discoideum 11 Figure 4. Immunofluorescent staining of DdCAD-1 in Ax2 and cadA null cells 29 Figure 5. Effect of RasG(G12T) expression from the rnrB promoter on discoidin mRNA 34 Figure 6. Discoidin expression in rasG null cells during axenic growth 36 Figure 7. Folate-repression of Discoidin protein levels in Ax2 and rasG null cells 37 Figure 8. Discoidin expression in rasG null cells during growth on bacteria 39 Figure 9. Discoidin expression during the development of rasG null cells 40 Figure 10. Induction of RasG(G12T) protein levels from the tet promoter and its effect on discoidin mRNA levels 42 Figure 11. Effect of RasG(G12T) expression from the tet promoter on the phosphorylation of vegetative proteins 44 Figure 12. Immunoblots of vegetative phosphoproteins separated by 2D gel electrophoresis.... 45 Figure 13. Representative changes in phosphorylation between the Ax2::MB and Ax2: :MB-rasG(G12T) strains 47 Figure 14. Comparison of protein phosphorylation levels between the Ax2::MB and Ax2::rasG(G12T) strains 48 Figure 15. Brilliant blue G-colloidal stained 2D gels of vegetative Ax2::MB and Ax2: :MB-rasG(G12T) lysates 50 Figure 16. Total protein levels of three putative RasG-responsive phosphoproteins in a RasG(G12T) expressing and vector control strain 54 Figure 17. Examination of the gefK and parental control strains for the presence of RasGEF-R phosphoprotein 55 Figure 18. Examination of the cadK and Ax2 strains for the presence of DdCAD-1 phosphoprotein 56 Figure 19. Induction of cell-to-cell adhesion during vegetative growth in a RasG(G12T) expressing strain 59 Figure 20. Disruption of RasG(G12T) mediated cell-to-cell adhesion with inhibitors of DdCAD-1 adhesion 61 Figure 21. Immunofluorescent localization of DdCAD-1 62 Figure 22. Cell-to-cell cohesion in developing Ax2::MB and Ax2::MB-ra,sG(G12T) cells 64 Figure 23. Levels of DdCAD-1 phosphorylation during development 66 Figure 24. Comparison of DdCAD-1 localization during development in Ax2: :MB and Ax2::MB-rasG(G12T) cells 67 Figure 25. Cellular streaming in Ax2, rasG null and cadA null cells 70 Figure 26. Levels of total and phosphorylated DdCAD-1 in a rasG null and Ax2 control cells. 72 Figure 27. Representative phosphorylation change in vegetative P K B between Ax2: :MB and Ax2::MB-rasG(G12T) cells 74 Figure 28. Folate induced stimulation of P K B phosphorylation 75 Figure 29. P K B membrane localization in response to RasG(G12T) expression 78 Figure 30. Chemotaxis of Ax2: :MB and Ax2::MB-ra.s'G(G12T) cells in a spatial folate gradient 79 Figure 31. Detection of a novel 70 kDa protein phosphorylated in response to folate 82 Figure 32. Fractionation of the folate-stimulated phosphoprotein p70 by 2D gel electrophoresis 83 Figure 33. Fractionation of the folate-stimulated phosphoprotein p70 on a narrow-pl range 2D gel 85 Figure 34. Sequence alignment of Dictyostelium LrrA and C. elegans SUR-8 86 Figure 35. A test for the reaction of LrrA with the phosophospecific M A P K antibody used to detect p70 phosphorylation 88 Figure 36. Detection of p70 in folate-stimulated IrrA nulls cells 89 Figure 37. 2D gel analysis of an IrrA null strain for the presence of p70 91 Figure 38. VatA overexpression changes the phosphorylation of p70 93 Figure 39. Subcellular fractionation of VatA and p70 94 ix LIST O F ABBREVIATIONS 2D two-dimensional A C adenylyl cyclase aleA aimless gene - RasGEF homologues B S A bovine serum albumin cAMP adenosine 3',5'-cyclic monophosphate cAR cAMP receptor CHAPS 3-[3-cholamidopropyl)dimethylammonio]-l-propanesulfonate C M F conditioned media factor DdCAD-1 Dictyostelium discoideum calcium-dependent adhesion molecule-1 DMSO dimethyl sulfoxide DTT dithiothreitol E D T A disodium ethylenediamine tetraacetic EGF epidermal growth factor EGFR epidermal growth factor receptor E R K extracellular regulated protein kinase FITC fluorescein isothiocyanate GAP GTPase activating protein GDI guanine nucleotide dissociation inhibitor GDP guanosine 5 '-diphosphate GEF guanine nucleotide exchange factor GPCR G-protein coupled receptor GTP guanosine 5 '-triphosphate HEPES N-[2-hydroxyethyl]piperazine-N'-[2-]ethanesulfonic acid HL5 nutrient rich axenic growth media H P L C high pressure liquid chromatography hrs hours IEF isoelectric focusing kDa kilodalton KK2 potassium phosphate buffer L R R leucine-rich repeat M A P K mitogen activated protein kinase min minutes M M S methyl methanesulfonate MS mass spectrometry MS V murine leukaemia virus P A G E polyacrylamide gel electrophoresis PBS phosphate-buffer saline P D K phosphoinositide dependent kinase PEG polyethylene glycol PH pleckstrin homology pi isoelectric point PI3K phosphoinositide 3-kinase P K A protein kinase A P K B protein kinase B PSF pre-starvation factor PTB phosphotyrosyl binding PtdIns(3,4,5)P3 phosphatidylinositol 3,4,5-bisphosphate PTEN phosphatase and tensin homologue P T K protein tyrosine kinase PVDF polyvinylidene difluoride REMI restriction enzyme-mediated integration RIP Ras-interacting protein R T K receptor tyrosine kinase SDS sodium dodecyl sulfate sec seconds SH2 Src homology 2 Sos Son of sevenless t time TBS Tris-buffered saline T E M E D N , N , N ' , N '-tetramethyl-ethylenediamine A C K N O W L E D G E M E N T S I would like to acknowledge the work and the play of those that I have spent anywhere from hours to days to years alongside while working towards this degree. In a room, lodged away in the middle of Wesbrook, time has flown and folks have come and gone, to each of you, my salutations. Specifically, I would like to thank Meenal Khosla for her help and collaboration on the discoidin project and for teaching me how to keep amoebae happy. M y appreciation to James Lim for his insight on assays and what a dusty piece of lab equipment, long forgotten, might be useful for. Thanks to Helmut Kae for his ability to ask and answer the question: "what would happen i f we did this?" To Michelle Wyse, the last of a wandering Weeks flock, I raise my glass to your inquisitive thinking and drive to understand science. To Parvin, Zahara, Shiv, Rujun, and all my other colleagues, I appreciate your time and effort. To my supervisors, Gerry Weeks and George Spiegelman, I express gratefulness and joy. Gerry you taught me to organize my thoughts and how to design my endeavors. George you taught me how to solve problems and think critically. Together you have taught me what I desired to learn and opened my mind to the possibilities. xn Dedicated to my father Who taught a boy that knowledge is astounding Bought him a plane ticket to B. C. And never tires of science 1 INTRODUCTION 1.1 A historical introduction to Ras research This thesis is part of an endeavor to further understand the role that Ras proteins play in regulating cellular functions. Research on Ras can be traced back 40 years to a murine leukaemia virus (MSV) and Jennifer Harvey, who showed this virus to be capable of inducing sarcomas in newborn rodents (Harvey, 1964). Viruses similar to the Harvey (Ha-MSV) strain were soon discovered, including Kirsten-MSV (Kirsten and Mayer, 1967), but their molecular characterization would wait some years due to a self-imposed ban on molecular cloning that was lifted in 1977 (Malumbres and Barbacid, 2003). Nonetheless, during this time it was becoming clear that these viruses were recombinant and that their oncogenic properties were due to their possession of genomic rat sequences (Scolnick et al., 1973). This came as an enormous shock and interest quickly shifted to these cellular 'oncogenes'. In 1979, reports appeared that genomic D N A from human tumor cell lines could transform NIH-3T3 cells, revealing that cellular genetic elements were capable of inducing these transformations (Shih et al, 1979; Perucho et al, 1981). Charming Der made the definitive connection three years later (Der et al., 1982), revealing that the Harvey and Kirsten viral oncogenes, which were given the acronym ras for rat sarcoma, were the same as the cellular oncogenes that had been used to transform NIH-3T3 cells. The study of human H-ras and K-ras was born and shortly thereafter a third highly related ras oncogene, N-ras, was identified (for review see Barbacid, 1987). This pioneering work on tumor inducing viruses, which many had originally thought to be "irrelevant", has sparked an extensive research field directed towards better understanding how Ras proteins regulate cell function (Malumbres and Barbacid, 2003). 1.2 An overview of Ras biology Ras proteins are undoubtedly important: the numerous biological functions to which they have been linked (e.g. proliferation, transformation, differentiation and apoptosis) and the fact that 20-30% of human cancers involve Ras mutations (Bos, 1989; Shields et al, 2000) should leave no question of this. In addition, all eukaryotic organisms thus far studied contain at least one highly conserved Ras protein (Barbacid, 1987; Bourne et al, 1991). This widespread distribution has been an asset to the goal of understanding Ras function, with information being 1 garnered from a variety of organisms, such as yeasts (Saccharomyces cerevisiae and Schizosaccharomyces pombe), Drosophila, Caenorhabditis elegans, Dictyostelium discoideum and mammalian cells (Egan and Weinberg, 1993; Wassarman et al., 1995; Campbell et al., 1998; Sternberg and Han, 1998; Weeks and Spiegelman, 2003). In the early 1990s, a synthesis of observations from these diverse organisms indicated a role for Ras in the transmission of signals from the cell surface to the nucleus (Egan and Weinberg, 1993). Since this time, the known physiological functions of Ras proteins have grown and for mammalian H-, K - and N-Ras proteins can be summarized into the five following categories. Firstly, it is well established that Ras proteins are involved in the transmission of mitogenic signals and proliferation (Campbell et al., 1998). In fact, Ras has been linked to cell cycle progression potentially through cyclin DI (Pruitt and Charming, 2001). Secondly, Ras proteins play essential roles in development; for example, mammalian K-ras is essential for mouse embryogenesis (Johnson et al., 1997). Thirdly, Ras has been linked to apoptosis and senescence, a process that may occur through the activation of the proteins such as p27 K i p l , p21 C i p l , p l 9 A r f and p l 6 1 N K 4 a (Ehrhardt et al., 2002). Fourthly, H-Ras has been linked to the regulation of cell adhesion through effects on integrin activation (Hughes et al., 1997). Lastly, Ras proteins have been implicated in regulating the actin cytoskeleton potentially through effects on Rac activation (Burridge and Wennerberg, 2004). This section will briefly review our current understanding of how Ras is thought to function in these roles. 1.2.1 The Ras superfamily of proteins Ras proteins comprise one subfamily of a large superfamily of proteins referred to as the Ras superfamily (Takai et al., 2001). This superfamily already numbers over 100 members that are divided in five subfamilies based on sequence relatedness: Rho, Rab, Sarl/Arf, Ran and Ras. A l l the members of the superfamily are small (20-40 kDa), monomelic proteins that bind guanine nucleotides (resulting in them often being referred to as GTP-binding proteins). Each subfamily appears to regulate distinct functions within the cell (reviewed in Takai et al., 2001); Rho subfamily proteins regulate cytoskeletal organization and gene expression; Rab and Sarl/Arf subfamily members regulate intracellular vesicle trafficking; Ran subfamily proteins regulate nuclear cytoplasmic transport and microtubule organization during the cell cycle; and Ras proteins integrate cell surface signals to cellular processes like proliferation, transformation, 2 differentiation and apoptosis. A great deal has been learned about each of the five subfamilies (reviewed by Bourne et al. 1991; Takai et al, 2001), but the rest of this section will deal solely with the Ras subfamily. 1.2.2 The G T P / G D P switch A l l members of the Ras superfamily function as nucleotide-dependent molecular switches (Figure 1). This function relies on the ability of these proteins to switch between two conformations: an active GTP-bound form and an inactive GDP-bound form (Bourne et al., 1991). The switch has been most extensively studied for Ras subfamily proteins, where the modulation between these two conformations requires three types of regulatory proteins. The first are guanine nucleotide exchange factors (GEFs) that promote the formation of active Ras-GTP, the second are GTPase activating proteins (GAPs) that promote the hydrolysis of Ras-GTP to Ras-GDP and the last are guanine nucleotide dissociation inhibitors (GDIs) that keep Ras in the GDP-bound form (Boguski and McCormick, 1993; Geyer and Wittinghdfer, 1997). As depicted in Figure 1, the concerted action of these proteins can be considered a cycle, which regulates the "on" (Ras-GTP) and " o f f (Ras-GDP) states of Ras, ultimately allowing Ras to act as a switch in signal transduction pathways. The importance of the Ras cycle is emphasized by the perturbations that occur as a result of various mutations of Ras. Single amino acid substitution at position 12 or 61 results in the locking of Ras in the active GTP-bound form, while substitution at position 17 prevents GTP binding and thus Ras activation (Barbacid, 1987). These mutations prevent normal Ras function by interfering with the GDP/GTP cycle. In fact, mutations that lock Ras in the active GTP-bound form are often found in human tumor cells (Bos, 1989). 1.2.3 Ras activation In mammalian cells, Ras is activated, i.e. stimulated to switch GDP for GTP, in response to the stimulation of receptor tyrosine kinases (RTKs), receptor-associated protein tyrosine kinases (PTKs) or G-protein coupled receptors (GPCRs) (reviewed in Schlessinger, 2000; Genot and Cantrell, 2000; Marinissen and Gutkind, 2001). Epidermal growth factor (EGF) mediated activation of Ras provides an example of the process with regard to R T K stimulation (Schlessinger, 2000; Carpenter, 2000). EGF stimulation of the EGF receptor (EGFR) promotes the autophosphorylation of various tyrosine residues in 3 Receptor activation Ras-GDP G E F Ras-GTP GAP v V V Rafl PI3K RalGDS Figure 1. A model of the Ras GTP/GDP switch. Cell receptor stimulation results in a guanine nucleotide exchange factor (GEF) facilitating the displacement of GDP and the binding of GTP to Ras. Ras-GTP then interacts and activates downstream effector molecules, such as Rafl , phosphatidylinositide 3-kinase (PI3K) and RalGDS. A GTPase activating protein (GAP) promotes the hydrolysis of Ras-GTP to RasGDP terminating Ras activity. 4 the cytoplasmic domain of EGFR. This tyrosine phosphorylation leads to the association of the adaptor proteins She and Grb2 through their Src homology 2 (SH2) and/or phosphotyrosyl binding (PTB) domains (Pawson and Scott, 1997). Grb2 and a RasGEF, Son of sevenless (Sos), then associate through an interaction between the C-terminus of Sos and the SH3 domains of Grb2 (Pawson and Schlessinger, 1993), bringing Sos into close proximity with Ras at the plasma membrane and allowing Ras activation through GDP/GTP exchange (Downward, 1996). Ras activation can also occur through the stimulation of receptor-associated PTKs, as in the case of ligand binding to immunoreceptors in B- and T-lymphocytes (Genot and Cantrell, 2000). In contrast to RTKs, these receptors possess no intrinsic tyrosine kinase activity and instead recruit PTKs to the membrane. Once recruited upon receptor stimulation, these PTKs phosphorylate tyrosine residues on immunoreceptors, which serve to further recruit Syk/Zap70 and adaptor proteins such as L A T and She (Genot and Cantrell, 2000). This results in the recruitment of the Grb2/Sos complex and the activation of Ras. The mechanism of Ras activation upon GPCRs stimulation has remained more elusive, but is believed to involve the GB? subunits of heterotrimeric G-protein and Src kinase acting to allow the transactivation of RTKs and thus Ras activation (Marinissen and Gutkind, 2001). The above paradigms represent an over-simplification and in reality other components also regulate the activation process. Thus, there are additional RasGEFs in mammalian cells, for example the Ras-guanine-nucleotide-releasing factor 1 (Ras-GRFl; also known as C D C 2 5 M m ) and closely related protein Ras-GRF2 that act as Ca2+-regulated Ras exchange factors (Cullen and Lockyer, 2002), the Ras guanine nucleotide-releasing protein (Ras-GRP; also known as CalDAG-GEFII) that acts as Ca 2 + - and diacylglycerol-regulated Ras exchange factors (Cullen and Lockyer, 2002), and three RasGEFs (MR-GEF, PDZ-GEF and GRP3) (Rebhun et al, 2000). Furthermore, 173 human genes have been identified whose encoded proteins are related to Ras superfamily GAPs and it has been suggested that at least 14 of these genes are RasGAPs (Bernards, 2003). 1.2.4 Ras Effectors Activated Ras interacts with a variety of effector proteins to transmit the activation signal downstream, a process that occurs through multiple cascades of cytoplasmic proteins (Marshall, 1996). Ras effectors preferentially interact with active Ras-GTP, as opposed to Ras-5 GDP, through the Switch 1 and Switch 2 regions of Ras, with the Switch 1 comprising amino acids 30-38 (which includes the Ras effector domain) and Switch 2 comprising amino acids 60-76 (Milburn et al, 1990; Marshall, 1996; Boriack-Sjodin et al, 1998; Herrmann, 2003). The Ras effectors bear little sequence homology but contain a structurally similar Ras binding domain (RBD) that is responsible for the interaction with Ras (Herrmann, 2003). Kinases of the Raf family, phosphoinositide 3-kinase (PI3Ks) and RalGEFs are three classes of proteins that have been sufficiently investigated to be considered as definite Ras effectors (Marshall, 1996; Campbell and Der, 2004). Other proteins have been implicated as Ras effectors, although their involvement is currently less well defined (Campbell et al., 1998; Cullen, 2001). Therefore, Ras effectors can be generalized into three groups: kinases, GEFs for other Ras Superfamily proteins and effectors with uncertain functions (Figure 2) (Ehrhardt et al., 2002). The first Ras effector to be identified was the serine/threonine kinase Raf (Vojtek et al., 1993). Shown to be recruited to the plasma membrane upon the activation of Ras and subsequently itself activated, the Raf kinase is an important transducer of biological signals (Morrison and Cutler, 1997; Takai et al., 2001). A cascade of cytoplasmic protein phosphorylation results from Ras activation of Raf (Morrison and Cutler, 1997), which begins with Raf phosphorylating and activating M E K , which in turn, phosphorylates and activates ERKs. Further transmission of the signal is achieved by the translocation of ERKs into the nucleus where they activate several transcription factors (e.g. Jun and Elk-1). Thus, the coupling of Ras to gene expression is mediated by the activation of kinase cascades that stimulate the phosphorylation and activation of transcription factors. The next-best-characterized Ras effector is phosphoinositide 3-kinase (PI3K) (Rodriguez-Viciana et al., 1997), which when activated in mammalian cells regulates apoptosis and rearrangements of the actin cytoskeleton (Shields et al., 2000). Activated PI3K catalyzes the formation of phosphatidylinositol 3,4,5-bisphosphate (PtdIns(3,4,5)P3) (Katso et al., 2001). PtdIns(3,4,5)P3 acts as a membrane docking site for the recruitment of pleckstrin homology (PH) domain containing proteins, such as PKB, also known as A K T (Cozier et al., 2004). The presence of PtdIns(3,4,5)P3 in the membrane causes P K B and phosphoinositide dependent kinase 1 (PDK1) to localize to the membrane. PDK1 then directly phosphorylates one of two sites on P K B that are required for P K B activation (Fresno Vara et al., 2004). The kinase responsible for 6 p r o l i f e r a t i o n p~ p r o l i f e r a t i o n , v e s i c l e t r a f f i c k i n g s u r v i v a l a c t i n c y t o s k e l e t o n g e n e e x p r e s s i o n • a p o p t o s i s • d i f f e r e n t a t i o n • e n d o c y t o s i s Figure 2. Downstream effectors of Ras proteins and their functions. Ras-GTP interacts with numerous effectors to regulate several cellular functions. Some Ras/effectors interactions, downstream components and subsequent cellular response are well characterized (top boxes), while others have yet to be critically tested or are unknown (bottom boxes). Adapted from Ehrhardt et al, 2002. 7 phosphorylating the second site is still unidentified, but it has been speculated that integrin-linked kinase (ILK) may be responsible (Persad et al., 2001). In mammalian cells, activated P K B then phosphorylates various downstream targets, including the apoptotic protein B A D , forkhead transcription factors and glycogen synthase kinase 3 (Cross et al., 1995; Datta et al., 1997; Biggs et al., 1999). In addition, Ras activation of PI3K can lead to the activation of Sosl/2, Vav, and Tiam-1, which are all capable of activating Rac, thereby regulating rearrangements of the actin cytoskeleton (Ehrhardt et al., 2002). Four RalGEFs (RalGDS, Rgl, Rlf, Rgr) have been described that also interact with Ras-GTP and subsequently function in the promotion of Ral activation (Wolthuis and Bos, 1999). Initially discovered as Ras interacting proteins in yeast two-hybrid assays (Hofer et al., 1994; Peterson et al., 1996), these proteins have since been confirmed to be in vivo effectors of Ras. Two additional RalGEFs, RalGPS/RalGEF2 and Rsc, have recently been described, but whether they interact with Ras proteins is not yet clear (Ehrhardt et al., 2002). Rapid activation of endogenous Ral is observed after the stimulation of a variety of receptors (e.g. receptor-associated tyrosine kinases and GPCRs) (Feig, 2003). The downstream consequence of this activation is less well defined, but some putative Ral targets have been described, including phospholipase D, RalBPl and Cdc42 (Wolthuis and Bos, 1999). Additional molecules have been described but still need to be critically tested as physiologically relevant Ras effectors (Campbell et al., 1998; Cullen, 2001; Ehrhardt et al., 2002). For example, phosphoinositide-specific phospholipase Ce (PLCe) was recently identified as a novel Ras effector (reviewed in Cullen, 2001), generating renewed speculation that Ras may be linked to the formation of diacylglycerol and inositol 1,4,5-triphosphate, which are known to regulate protein kinase C (PKC) and C a 2 + signaling respectively (Williams, 1999). Another example is R i n l , a Ras subfamily protein that can form a stable complex with Ras in vivo (Campbell et al, 1998) and has been suggested to facilitate Ras-mediated endocytosis (Tall et al., 2001). The serine/threonine kinase M E K K 1 has been shown to bind directly to the effector domain of Ras (Russell et al., 1995) and be an upstream activator of the Jun N-terminal kinase cascade (Campbell et ah, 1998). Examples of other candidate effectors include AF-6, calmodulin and Nore-1 (Ehrhardt et al., 2002). Through these, it has been suggested that Ras regulates cell-cell junctions, Ca signaling and apoptosis respectively (Campbell et al, 1998; Cullen, 2001; 8 Ehrhardt et al, 2002). This growing number of Ras effectors reveals that much is still to be learned. 1.3 A broader understanding of Ras function through model organisms Much of the information described above has been obtained using in vitro approaches and transfected mammalian cell lines. However, the conservation of Ras proteins throughout eukaryotes provides the potential to broaden of our understanding of Ras function. In fact, studies with the model organisms C. elegans and Drosophila have already made a significant impact on our understanding of Ras. For example, the C. elegans Ras homologue Let60 has been linked to the formation of the vulva during larval development (Sternberg and Han, 1998) and Drosophila Rasl activation is associated with the differentiation of the R7 photoreceptor cell during development of the compound eye (Wassarman et al., 1995). In addition, studies in Dictyostelium have uncovered an unusually large Ras subfamily of proteins, several of which have already been genetically disrupted, uncovering novel Ras functions (Weeks and Spiegelman, 2003). Thus, the conservation of signaling strategies throughout eukaryotic evolution has been particularly useful in furthering our understanding of the role of Ras in signal transduction. 1.4 Dictyostelium as a model organism Dictyostelium has proved to be an excellent system for the study of signaling networks and parallel pathways (Parent and Devreotes, 1996). Many of the proteins involved in mammalian signaling (e.g. GSK-3 and STAT) are conserved in Dictyostelium (Harwood et al., 1995; Kawata et al., 1997) but not in other unicellular organisms like yeast. In addition, the study of these proteins is greatly facilitated by the availability of a genetic approach that allows the analysis of novel genes and pathways (Parent and Devreotes, 1996; Guerin and Larochelle, 2002). This includes high-efficiency transformation with extrachromosomal and integrating vectors, the ability to disrupt or replace genes by homologous recombination, and insertional mutagenesis using restriction enzyme-mediated integration (REMI). The near completion of the Dictyostelium genome is quickening the pace of this research through the further identification of unique and conserved genes (Eichinger and Noegel, 2003). In fact, the Dictyostelium genome sequence has already predicted at least five new and as yet uncharacterized ras genes (M. Wyse, unpublished observations). 9 One of the appeals of Dictyostelium as a model organism is its life cycle, which allows growth and differentiation to be studied as separate processes (reviewed in Kessin, 2001). This solitary amoeba generally feeds on bacteria during its vegetative phase, or in case of axenically created laboratory strains ingests liquid media. Upon nutrient deprivation, Dictyostelium initiates a developmental program where free-living amoebae chemotax towards cAMP and aggregate into a multicellular organism (Figure 3). Within the aggregate different cell-types (prespore and prestalk) begin to emerge and spatially organize. Prestalk cells sort to the top of the aggregate where a single tip forms and the structure subsequently elongates to produce a migrating slug or pseudoplasmodium. Later, culmination results in the formation of a mature fruiting body, in which prestalk cells produce a fibrous stalk that supports a prespore-derived spore mass. The entire process is rapid, occurring within 24 hours. cAMP is a key chemoattractant during Dictyostelium aggregation, and signaling pathways that regulate this process have been well documented (reviewed in Aubry and Firtel, 1999). c A R l , a seven transmembrane glycoprotein belonging to the GPCR superfamily, is central to this process (Kessin, 2001). c A R l is one of a group of four receptors (the others are cAR2, 3 and 4) whose ligand is cAMP. These receptors are associated with heterotrimeric G-proteins. The genes for eight Ga, one G3 and one Gy protein, capable of forming these complexes, have been cloned (Kessin, 2001; Zhang et al., 2001). The binding of cAMP to c A R l results in adenylyl cyclase A (ACA) activation, the transcriptional activation of aggregation specific genes and the mobilization of the cytoskeleton (e.g. the accumulation of actin and myosin II in the cytoskeleton) (Aubry and Firtel, 1999). The activation of A C A is a requirement for the relay of the cAMP signal during aggregation and requires the release of the GBy subunit from c A R l upon reception of a cAMP signal. At least two additional proteins are required for A C A activation: C R A C and PiaA (Insall et al., 1994; Chen et al., 1997). Mutations in genes encoding potential Ras pathways components: ERK2, AleA (a RasGEF), and RasC (discussed below), all cause defects in aggregation (Segall et al., 1995; Insall et al., 1996; Lim et al., 2001). Furthermore, a PI3K dependent pathway has been described that is activated in response to cAMP (Zhou et al., 1995), since cells lacking both PI3K1 and PI3K2 (pi3KF/pi3k7) show defects in chemotaxis and aggregation, and the PH-domain containing proteins P K B and PhdA require PI3K1/2 activity for their localization in response to cAMP (reviewed in Firtel and Chung, 2000). 10 Figure 3. Mul t ice l lu la r development of Dictyostelium discoideum. Dictyostelium discoideum developmental structures are shown in a clockwise pattern beginning with (a) an aggregating mound of cells, (b) standing slug or finger, (c) migrating slug, (d) 'mexican hat', (e) culminating structures, and if) a terminal fruiting body. The process from a to /takes approximately 24 hours to complete. Electron micrograph image by M . G. Grimson and R. L. Blanton, Texas Tech University, Lubbock, Texas. 11 At later stages of development, morphogens including differentiation-inducing-factor (DIF), adenosine, and ammonia, are important for cell-type decisions (Parent and Devreotes, 1996; Kay et al, 1999). DIF-1, the most extensively studied, induces stalk cell formation, while inhibiting prespore cell formation (Kessin, 2001). Several other molecules have also been shown to influence cell-type decisions, including glycogen synthetase kinase-3 (GSK-3), Wariai and M E K K a (Weeks, 2000). Extracellular cAMP, through cAR3, regulates the activity of GSK-3 and the protein tyrosine kinase ZAK1 has been implicated as an intermediate in this process (Plyte et al, 1999; Kim et al, 1999). Less is known about signaling in vegetative cells and the transition from growth to development, although a few processes have been examined in some detail. For example, growing cells chemotax towards bacteria, attracted by secreted chemoattractants including folic acid and pterin (Kuwayama et al, 1993; Parent and Devreotes, 1996). In the chemotactic response to folate, the binding of folic acid to its unknown receptor results in the activation of guanylyl cyclases and alteration of the cytoskeleton (Kuwayama et al, 1993; Blusch and Nellen, 1994). In addition, growing cells secrete various factors, including pre-starvation factor (PSF) and conditioned media factor (CMF) (Clarke and Gomer, 1995). PSF is continually secreted by growing cells and triggers the expression of genes whose products are required for aggregation, although it is not known how this is mediated (Burdine and Clarke, 1995). C M F is secreted upon starvation, where it participates in regulating aggregation (Jain et al, 1992; Yuen et al, 1995). One pathway involved in the transition from growth to development that has been partially characterized involves the protein kinase YakA (Souza et al, 1998). yakA null cells are smaller, have a cell cycle that is accelerated, do not turn off genes expressed during growth when starved and do not initiate development (Souza et al, 1998). The use of REMI on yakA null cells revealed that the disruption of the gene encoding the translational regulator PufA reversed their developmental block (Souza et al, 1999). The disruption of pufA in yakA null cells restores the synthesis of P K A - C and PufA can bind to P K A - C , suggesting a pathway where PufA inhibits P K A - C synthesis and YakA inhibits PufA (Souza et al, 1999). In addition, YakA has recently been implicated in the folate-stimulated activation of guanylyl cyclases and alteration of the cytoskeleton (van Es et al, 2001). 12 1.5 Dictyostelium Ras proteins Six distinct Ras subfamily proteins, encoded by the genes rasG, rasD, rasB, rasS, rasC and rapA, have thus far been characterized in Dictyostelium (reviewed in Daniel et al., 1995). The five Ras proteins listed above are well conserved relative to the human H- , N - , and K-Ras proteins, sharing between 54 and 68% amino acid identity with H-Ras. The only Dictyostelium Rap protein thus far identified, encoded by rapA, is even more conserved, sharing 76% amino acid identity to human Rapl . Several novel ras genes have emerged from analysis of the Dictyostelium database, and although these Ras proteins are less well conserved, they are clearly members of the Ras subfamily (M. Wyse, personal communication). Investigation into the specific functions of the individual Ras proteins has been centered on ras gene ablation studies, but studies on the overexpression of activated or dominant negative forms of Dictyostelium Ras proteins have also provided insight. The first Dictyostelium Ras protein to be studied in this way was RasD. Transformants overexpressing an activated version of RasD, RasD(G12T), were found to form multi-tipped aggregates that were blocked in further development (Reymond et al., 1986). These transformants express enhanced levels of two prestalk cell-specific markers and decreased levels of a prespore cell-specific marker (Louis et al, 1997), suggesting that RasD influences cell fate. Activated Ras expression from a prespore promoter was sufficient to reproduce the multi-tipped phenotype (Jaffer et al., 2001), suggesting that activated protein exerts its influence in the prespore cell population. It was thus surprising when the only observable phenotype of a rasD null strain was defective phototaxis and thermotaxis of the slug (Wilkins et al, 2000b). However, it has been suggested that RasG, which share 82% overall amino acid identity and identical effector regions with RasD, might be compensating for RasD function in these cells (Weeks and Spiegelman, 2003). Aggregation was recently shown to require RasC, since rasC null cells are unable to produce cAMP upon receptor stimulation (Lim et al, 2001). Thus, although rasC null cells do not normally aggregate, this phenotype can be circumvented by exogenously applying cAMP to the cells. In addition to its requirement for adenylyl cyclase activation, RasC is also required for wild type levels of P K B activity (Lim et al, 2001; Lim, 2002). Starved rasC null cells also have motility defects and chemotax poorly to cAMP, but interestingly, cAMP-pulsed rasC null cells move more rapidly toward cAMP (Lim et al., 2001). Cells expressing an activated version of 13 RasC, RasC(G13T), have also been created and exhibit reduced motility and defects in aggregation (Lim, 2002). Disruption of rasS resulted in cells that were unable to grow in axenic culture due to impairments in fluid phase endocytosis (Chubb et al, 2000). rasS null cells also move rapidly, are highly polarized and exhibit increased F-actin localization to their pseudopods. Based on these observations it was suggested that RasS regulates the balance between feeding and movement, with competition for the cytoskeletal components required for both feeding and motility (Chubb et al, 2000). Thus, the rapid movement of rasS null cells would impact their ability to feed. A strain null for a RasGEF protein, RasGEF-B, also show reduced endocytosis and enhanced motility. While it was initially suggested that RasGEF-B regulates RasS (Wilkins et al., 2000a), the motility defects in rasS and gefB null cells have been shown to rely on different mechanisms (Chubb et al, 2002), even though rasS/gefB double null strain shows phenotypes similar to each single null strain (King and Insall, 2004). Both rasB and rapA appear to be essential for growth (Sutherland et al., 2001b; Kang et al., 2002). A l l presumptive rasB null cells grew very slowly but contained at least two copies of the disrupted gene and eventually reverted to a wild type phenotype (Sutherland et al., 2001b). RasB has been shown to be localized to the nucleus in cell cycle specific manner and RasB(G12T) expression has been shown to produce multinucleate cells (Sutherland, 2001a; Sutherland et al, 2001b). Attempts to disrupt the rapA have been unsuccessful. The reduction of Rapl levels by antisense reduces growth and viability, suggesting the rapA is an essential gene (Kang et al, 2002). Rapl is activated in response to osmotic stress (Kang et al, 2002). The most studied Dictyostelium Ras protein has been RasG. This gene was identified using low stringency hybridization with the only other Dictyostelium ras gene that had been identified at the time, rasD, as a probe (Robbins et al, 1989). RasG was subsequently found to be the major Ras species present in vegetative cells, with its expression down regulated to negligible levels by the aggregation stage (Robbins et al, 1989; Khosla et al, 1990). This suggested a role for RasG during growth, although, the overexpression of an activated form of RasG, RasG(G12T), and the creation of a rasG null strain indicated a possible role for the protein in early development (Khosla et al, 1996; Tuxworth et al, 1997). Cell expressing RasG(G12T) exhibit several phenotypes. These cells were unusually flattened with numerous filopodia and prominent dorsal and peripheral membrane ruffles that 14 contained large amounts of F-actin (Zhang et al, 1999), suggesting the possibility that RasG(G12T) expression affects the actin cytoskeleton. These cells also showed impaired motility and chemotaxis compared to wild type strains (Khosla et al, 1996). However, although the effects of RasG(G12T) expression on the cytoskeleton and motility were conspicuous, the most noteworthy phenotype of cells expressing RasG(G12T) was their inability to aggregate, with these cells producing less cAMP than a wild type strain in response to a pulse of 2?-deoxy-cAMP (Khosla et al., 1996). This last phenotype suggested that constitutive RasG signaling blocked the initiation of development and implicated RasG in potentially regulating the transition from growth to development. The rasG null cells also exhibited a complex phenotype. Vegetative rasG null cells displayed multiple elongated filopodia, aberrant lamellipodia and unusual punctate polymerized actin structures (Tuxworth et al, 1997). These cells lacked cell polarity and it was thus not surprising that they also exhibit both reduced motility and chemotaxis (Tuxworth et al., 1997; Khosla et al, 2000). The growth of these cells in shake suspension is very slow, and growth was to a much lower final density than parental control strains. Suspension grown cells were multinucleate, being defective in the completion of cytokinesis due to an inability to sever the cleavage furrow (Tuxworth et al., 1997). This cytokinesis defect was not observed in rasG null cells grown on plastic dishes, presumably due to the ability of Dictyostelium cells to overcome cytokinesis defects on surfaces by traction-mediated cytofission (Fukui et al., 1990). The loss of RasG appears to be partially compensated for by other Ras proteins, in that rasG null cells contained increased levels of RasD, RasB and RasC (Khosla et al., 2000; Weeks and Spiegelman, 2003). The first downstream signaling protein linked to RasG was ERK2, a M A P kinase activated in response to cAMP (Segall et al., 1995). Use of antibodies that react with phosphorylated ERK2 (Kosaka et al., 1997) revealed that cells expressing RasG(G12T) exhibited decreased ERK2 phosphorylation in response to cAMP (Aubry et al., 1997; Kosaka et al., 1998), whereas cells expressing a dominant negative rasG gene exhibited increased ERK2 phosphorylation (Aubry et al., 1997). RasG has also been shown to interact with Dictyostelium PI3K1 and PI3K2 in yeast two-hybrid assays (Lee at al, 1999; Funamoto et al, 2002). Cells lacking both PI3K1 and PI3K2 exhibit defects in aggregation and chemotaxis (Buczynski et al, 1997; Funamoto et al, 2001) 15 and these proteins are required for the translocation PH-domain containing proteins to the membrane (Meili et al, 1999; Funamoto et al, 2001). In addition, PI3K activity, which results in the generation of PtdIns(3,4,5)P3 and PtdIns(3,4)P2 (Katso et al, 2001), has been shown to be activated by chemoattractant stimulation and confined to the leading edge of chemotaxing cells (Huang et al, 2003). Since RasG binds to PI3Ks, it could well play a role in regulating these processes. Ras-interacting protein 3 (RIP3) was also identified in a yeast two-hybrid assay as a putative RasG interacting protein (Lee at al, 1999). rip3 null cells are unable to aggregate or activate adenylyl cyclase upon receptor activation (Lee at al, 1999). Unfortunately, RIP3 lacks a recognizable homolog in other organims (Lee at al, 1999) and it is not clear as yet how RasG binding to RIP3 affects its function. Nonetheless, the identification of this protein suggests a potential role for RasG in regulating aggregation and receptor-stimulated adenylyl cyclase activity. 1.6 Thesis Objectives 1.6.1 Background Following on from studies on the expression of RasG(G12T) under the control of the folate-repressible discoidin promoter in apVEII-rasG{G\2T) transformant (Khosla et al, 1996), it was found that levels of RasG(G12T) increased as growth progressed in the absence of folate. However, the expression of RasG(G12T) from the discoidin promoter reached a maximum at a cell density of approximately 7 x 10s cells/ml and then declined dramatically (Secko et al, 2001). Discoidin expression is known to decrease at high cell density (Wetterauer et al, 1995), but the observed decline in RasG(G12T) expression (Secko et al, 2001) was more dramatic and occurred at a considerably lower cell density than had been previously reported. A possible explanation for the difference was that the elevated level of RasG(G12T) dramatically inhibited expression from the discoidin promoter. To further investigate the possible link between RasG(G12T) levels and discoidin expression, the levels of Discoidin in the pVEII-rasG(G12T) transformant was compared with those in the Ax2 parental strain at various stages during growth after the removal of folate. As cell density increased there was the anticipated increase in Discoidin in response to the removal of folate for both the parental Ax2 cells and the pVEII-rasG(G\2T) transformant and this 16 increase occurred at similar cell densities in the two strains. However, considerably more Discoidin was produced in the Ax2 parental cells than in the pVEII-rasG(G\2T) transformant and the levels of Discoidin in the pVEII-rasG(G\2T) transformant declined at a lower cell density as growth progressed. These results are consistent with the idea that elevated levels of RasG(G12T) repressed discoidin expression. However, since RasG(G12T) was expressed on the discoidin promoter, it was difficult to definitively interpret the results. 1.6.2 Objectives The first objective of this thesis was to confirm that activated RasG inhibited discoidin expression. This required the development of an alternative inducible expression system, which was accomplished through the use of the inducible rnrB promoter. The second objective was use an expression system to rapidly express activated RasG to allow the consequences of the RasG activation signal to be monitored. The development of a system to rapidly express activated RasG provided a method to investigate the virtually unknown RasG signaling pathway and this investigation comprised the third objective of this thesis. This objective relied on Ras proteins making extensive use of phosphorylation cascades involving multiple kinases and phosphatases in many other organisms (see Section 1.2; Campbell et al., 1998; Shields et al., 2000). Thus, the third objective was to monitor the changes in the phosphorylation of proteins that occurred upon RasG(G12T) expression and to identify these proteins using proteomic techniques. 17 2 M A T E R I A L S A N D M E T H O D S 2.1 Materials 2.1.1 Reagents Chemicals were acquired from Sigma, Fisher Scientific, ICN, Bio-Rad, B D H or Invitrogen, unless otherwise stated. Restriction enzymes, D N A polymerases and other D N A modifying enzymes were obtained from New England Biolabs or Invitrogen. Growth media reagents were from Oxoid, B B L , or Difco. Chemical or electro-competent Escherichia coli strain X L l - B l u e M R F ' (Stratagene) was used for all cloning and sub-cloning procedures. Immobiline DryStrips, PVDF membranes, enhanced chemiluminescence assay and secondary antibodies were from Amersham Pharmacia Biotech. Blasticidin S was from Calbiochem. Modified sequencing grade trypsin and complete protease inhibitor cocktail were from Roche Diagnostics. Siliconized microcentrifuge tubes were from Diamed. 40% Acrylamide/Bis Solution (19:1, electrophoresis grade) and Dc Protein Assay were from Bio-Rad Laboratories. Reagents used for 2D electrophoresis and mass spectroscopy (MS) analysis (except acrylamide/bis solution) were of HPLC quality. Phospho-p44/42 M A P kinase polyclonal antibody (Cat # 9101), phosphothreonine polyclonal antibody (Cat # 9381), phosphotyrosine monoclonal antibody (Cat #9419), Myc-tag polyclonal antibody (Cat # 2272) were from Cell Signaling Technology. Monoclonal actin-specific antibody was from Chemicon (Cat # MAB1501). The Dictyostelium specific antibodies were generously provided as follows: the PKB-specific polyclonal antibody by Dr. Robert Dottin (Hunter College, New York), the DdCAD-1-specific polyclonal antibody by Dr. Chi-Hung Siu (University of Toronto, Toronto), the Dictyostelium VatA-specific N4 antibody and the Discoidin I antibody by Dr. Margaret Clarke (Oklahoma Medical Research Foundation, Oklahoma City). The RasG and RasC specific antibodies were generated in the Weeks laboratory (Khosla et al, 1994; Lim, 2002). 2.1.2 Services D N A sequencing was performed at the University of British Columbia Nucleic Acid and Protein Services (NAPs) unit (www.biotech.ubc.ca/services/naps/index.html). 18 Oligonucleotides for PCR were generated at Alpha D N A (www.alphadna.com). Mass spectral analysis was performed at the UVic-Genome B C Proteomics Centre (www.proteincentre.com). Indirect immunofluorescence was performed at the University of British Columbia Biolmaging Facility (www.emlab.ubc.ca). 2.1.3 Dictyostelium strains Ax2 is the parental strain used for all transformations and this strain was originally obtained from Dr. Barry Coukell (York University, Toronto). Ax2 is an axenic derivative of the wild type strain NC-4 (for a discussion of strain history please see: dictybase.org/strain_history.htm). The pVEII-rasG(G\TT) strain was generated in the Weeks Laboratory (Khosla et al., 1996). The rasG null and gefR null strains were obtained from Dr. Robert Insall (University of Birmingham, England). The cadA null strain was obtained from Dr. Chi-Hung Siu (University of Toronto, Toronto). The IrrA null strain was obtained from Dr. Wen-Tsan Chang (National Cheng Kung University Medical College, Taiwan). The vatA-oe strain was obtained from Dr. Margaret Clarke (Oklahoma Medical Research Foundation, Oklahoma City). 2.1.4 Plasmids Plasmids were obtained from the indicated sources: the tetracycline expression vectors, MB-35 and MB-38, from M . Blaauw (University of Groningen, Groningen), ptz-rasG(G12T) from Meenal Khosla (UBC), RnrB-ubi-S65TGFP6 from Dr. Harry MacWilliams (Ludwig-Maximilians-Universitat, Munich), discoidin Iy gene fragment from Dr. Margaret Clarke (Oklahoma Medical Research Foundation, Oklahoma City). 2.2 M e t h o d s 2.2.1 Dictyostelium cell culture Dictyostelium discoideum cells were grown axenically in HL5 medium (14.3 g peptone, 7.15 g yeast extract, 15.4 g glucose, 0.96 g Na 2 HP0 4 -7H 2 0 and 0.486 g K H 2 P 0 4 per liter of water) supplemented with 50 ug/ml streptomycin (Sigma) at 22°C, either on tissue culture dishes (Nunc) or in rotatory agitated suspension (175 rpm) (Watts and Ashworth, 1970). Transformed strains were selected and maintained in HL5 media supplemented with the either: 5 ug/ml 19 blasticidin S (Calbiochem) or 10 pg/ml G418 (Gibco-BRL) as appropriate. rasG null strains were grown and maintained on Nunc tissue culture dishes, since this strain grows poorly in rotatory agitated suspension (Tuxworth et al, 1997), unless otherwise stated. Ax2: :MB, AX2::MB-rasG(G12T) and A x 2 : : M B - L r r A M Y C strains were additionally maintained and grown in 5 pg/ml tetracycline (Sigma) until their use and thepVEII-rasG(G\2T) strain was maintained in 1 m M folate until required. Cells grown in shake suspension were passaged by diluting between 20-50 fold in fresh media upon reaching a cell density of approximately 2-3 x 106 cells/ml. Cells numbers were determined using a hemocytometer. Cells grown on Nunc plates were passaged by diluting 10-30 fold into fresh media/plates when they research confluent density. A l l cells were maintained for no more than 7 passages before being re-derived from cryo-preserved stocks or spores. For cryo-preservation, approximately 2.5 x 107 cells/ml in 1.0 ml aliquots were frozen at -70°C in HL5 containing 10% DMSO. 2.2.2 Dictyostelium growth in bacterial suspension Axenically grown cells were inoculated into a suspension of Klebsiella oxytoca in K K 2 , OD6oo of 8 (Rathi and Clarke, 1992; Primpke et al, 2000) and allowed to grow for approximately seven to eight generations before being transferred to a fresh suspension of bacteria. Dictyostelium cells were collected by centrifugation at 500 x g for 3 minutes, then washed exhaustively to remove residual bacteria before being used for further analysis. 2.2.3 Dictyostelium development Vegetative cells were axenically grown to a density of 2-3 x 106 cells/ml in shake suspension or to confluence on Nunc plates, washed twice in 20 m M potassium phosphate buffer, pH 6.5 (KK2), and resuspended in K K 2 to a density of 5.0 x 106 cells/ml. Cells were then shaken at 160 rpm for the indicated periods at 22°C. Cells were also grown in association with bacteria to a density of approximately 2-3 x 106 cells/ml, exhaustively washed with K K 2 to remove bacteria, plated at 1 x 106 cells/cm2 on non-nutrient agar (2% agar buffered in K K 2 , ImM MgCl2, 0.1 m M CaCl 2) and allowed to develop for the indicated periods. 2.2.4 Vector constructions and Polymerase Chain Reaction (PCR) To generate the rnrB-rasG{G\2T) construct, a vector containing the rnrB promoter, RnrB-ubi-S65TGFP6 (Gaudet et al, 2001), was digested with Xhol to remove gfp, treated with 20 Klenow polymerase and deoxynucleoside triphosphates to generate blunt ends and then subsequently digested with BgRl. A rasG(G12T) insert (Khosla et al, 1996) was then sequentially digested with BgRl and Smal and ligated to the above vector fragment. To generate the MB-rasG(G12T) vector, the tetracycline responsive vector MB-38 (Blaauw et al, 2000) and the rasG(G12T) insert described above (Khosla et al, 1996) were both digested with BgRl and Mlul and ligated together. The MB-rasG(G12T) and rnrB-ras G(G12T) constructs were transformed into Escherichia coli strain X L 1 - M R F (Stratagene) and their sequences confirmed by D N A sequencing. The IrrA (# SLG759) and vatA (# VHD171) cDNAs were obtained from the Japanese Dictyostelium cDNA project (http://www.csm.biol.tsukuba.ac.jp/cDNAproject. html). The primer l r rAFl (5' - G A A T T C A T G G G A G G A A A T T T A T C A T - 3') and IrrARmyc (5' -T T A A A G A T C C T C T T C T G A A A T A A G T T T T T G T T C T G C T G C T T T T T C A T T A T C T T G T T G A A T A C C - 3') were used for PCR to amplify the IrrA cDNA while adding the c-myc epitope EQKLISEEDL (Lim, 2002) before the termination codon of the IrrA cDNA. The primers vatFl (5' - G A G C T C T G C A C G A G A A A T G T C A A A A A A T T C - 3 ' ) and vatRmyc (5' - T C T A G A T T A A A G A T C C T C T T C T G A A A T A A G T T T T T G T T C T G C T G C A A C T A A A T C A C T A A A G T T T C - 3') were used for PCR to amplify the vatA cDNA, adding the same c-myc epitope as above. A l l PCR reactions were performed using an Idaho RapidCycler (Idaho Technologies). The quantities of template D N A used per reaction were 0.1 - 0.5 ng plasmid D N A . A standard PCR reaction to test primers was done in 10 ul mixtures in glass capillary tubes containing 5.38 ul ddH 2 0, 1.0 ul lOx Idaho PCR buffer (500 m M Tris, pH 8.3, 2.5 mg/ml BSA, 20% (w/v) sucrose, 1 m M cresol red, 30 m M MgCl 2 ) , 0.5 u\ 2.5 m M dNTPs (Pharmacia), 1.0 ul of each 2.5 uM oligonucleotide, 1.0 ng D N A and 0.125 ul Taq D N A polymerase (5 U / al) (Gibco-BRL). High fidelity PCR with the proof reading enzyme Pfu Turbo D N A polymerase (Stratagene) was used to construct all expression vectors after testing the primers. High fidelity PCR reactions were done in 10 /xl mixtures in plastic capillary tubes containing 4.7 ng ddH 2 0, 1.0 ul lOx Cloned Pfu D N A polymerase reaction buffer, 1.0 al 2.5 m M dNTPs (Pharmacia), 1.0 ul of each 2.5 uM oligonucleotide, 1.0 al D N A and 0.3 u\ Pfu D N A polymerase (2.5 U / ul). For both standard and high fidelity PCR the cycling parameters were as follows: (i) 2 cycles of D = 92°C for 60 sec, A = x°C for 7 sec, E = 72°C for 90 sec; (ii) 36 cycles (Taq) or 30 cycles (Pfu) ofD = 92°C for 1 sec, A = x°C for 7 sec, E = 72°C for;; sec; hold at 72°C for 120 21 sec. (Where D is denaturation, A is annealing, E is extension, x is the temperature optimized for a particular pair of primers, and y is 60 sec for every 1000 bp of amplification target.) The temperature ramping rate for all cycling was set at 6.0. Amplified PCR fragments for IrrA and vatA were ligated into pGEM-T Easy (Promega) to generate the vectors pGEM-lrrAmyc and pGEM-vatAmyc. These vectors were subsequently digested with Sphl and Mlu\ and ligated into the SphVMlul digested MB-38 vector to generate the vectors MB-lrrAmyc and MB-vatAmyc. 2.2.5 Dictyostelium transformations To generate the tetracycline-responsive Ax2: :MB, AX2::MB-rasG(G12T) and A x 2 : : M B - L r r A M Y C strains, the MB38, MB38-rasG(G12T), MB3S-lrrAmyc and MB35 vectors were first purified by PEG precipitation (Sambrook et al., 1989). Ten pg of the MB38, MB38-rasG(G\2T) and MB38-/rr^4mvc vectors were then mixed with ten pg of MB-35 and incubated with 1 x 107 Ax2 cells that had been previously washed and resuspended in 0.8 ml electroporation buffer (10 m M NaP04, pH 6.1, 50 m M sucrose). This suspension was incubated on ice for 10 min, pulsed with a Bio-Rad GenePulser set at 0.9 kV, 3 uF, and then incubated on ice for 10 min. HL5 media supplemented with 5 pg/ml tetracycline was added and the cells were plated in Nunc tissue culture dishes at 22°C. 24 hours later the media was replaced with fresh HL5 media supplemented with 5 pg/ml blasticidin S, 10 pg/ml G418 and 5 pg/ml tetracycline. Transformants were grown for 10 days, before stable transformants were selected by clonal growth in HL5 containing G418, blasticidin S and tetracycline at the previously indicated concentrations. The rnrB-rasG(G\2T) transformant was obtained by the CaP04 D N A precipitation method (Nellen et al., 1987). This involved precipitating 10 pg of plasmid D N A for 0.5 hr in a mixture containing 375 pi ddH 2 0, 125 pi 0.125 M CaCl 2 and 500 pi 2X HBS (42 m M HEPES, 270 m M NaCl, 10 m M K C L , 1.4 m M Na 2 HP0 4 , 10 m M glucose, pH 6.1). Precipitates were then added drop wise to Dictyostelium cells cultured in Nunc dishes in Bis-Tris buffered HL5 (2.1 g Bis-Tris, 10 g peptone, 5 g yeast extract, 10 g glucose per 1 liter of dH 2 0) and incubated at 22°C for 4 hr. Cells were then exposed to a solution of 15% glycerol/lX HBS for 2 min, followed by replacement of HL5 media. 24 hours later this media was replaced with HL5 medium supplemented with 10 pg/ml of G418. Colonies were visible after 10-14 days and stable 22 transforrnants were selected by clonal growth in HL5 medium containing G418 and streptomycin at the previously indicated concentrations and were maintained in the same medium. 2.2.6 Induction conditions for protein overexpression ApVEII-rasG(G12T) cell population was centrifuged at 700 x g for 4 min, washed once in HL5 media and resuspended in HL5 media containing no folate. The rnrB-ras G(G\2T) strain was treated as above and then incubated with 5 or 10 m M methyl methanesulfonate (MMS) (Sigma) to induce protein expression. A l l tetracycline repressible strains were centrifuged as above, washed three times in HL5 media, and then resuspended in HL5 media containing no tetracycline to induce protein expression. For all strains, incubation was performed in rotatory agitated suspension (175 rpm) at 22°C. 2.2.7 RNA analysis Total cytoplasmic R N A was isolated from 1 x 107 cells using a modified protocol employing the TRIzol Reagent (Gibco-BRL). Pelleted cells were lysed in 1.0 ml TRIzol and incubated for 5 mins at 22°C, then mixed with 200 ul chloroform and incubated a further 5 min. Sample were centrifuged at 16,000 x g for 15 min and the upper aqueous phase (~ 600 ul) was then transferred to a new microcentrifuge tube. The upper aqueous phase was mixed with 500 ul of iso-propanol, incubated 10 min and centrifuged at 16,000 x g for 15 min. The pelleted R N A was further washed with 1.0 ml 75% ethanol, air dried, dissolved in DEPC-treated ddH 2 0. Dissolved R N A was quantitated by absorbance spectrophotometry at 260 nm. Samples were size fractionated by electrophoresis on 1% formaldehyde-agarose gels as previously described (Khosla et al, 1996). Equal loading of samples was verified by observing the intensities of 28S and 18S rRNAs following ethidium bromide staining. The R N A was transferred to a nylon membrane (Hybond N+, Amersham) and the filters were prehybridized for 3 to 4 h at 42°C in a mixture containing 30% formamide, 5x Denhardt's solution ( lx Denhardt's solution is 0.02% Ficoll, 0.02% bovine serum albumin, and 0.02% polyvinylpyrrolidone), 5x SSC (lx SSC is 0.15 M NaCl plus 0.015 M sodium citrate), 20 m M sodium phosphate buffer (pH 6.5), 0.5 % sodium dodecyl sulfate (SDS), and 30 ixg of polyadenylic acid per ml. A 330 bp Hindlll/EcoRl discoidin ly gene fragment (Rathi et al., 1991) that had been first gel purified using the QIAquick Gel Extraction Kit (Qiagen) was labeled with [a- 3 2P]dCTP by the random primer method as 23 previously described (Sambrook et al., 1989). This probe was added to the prehybidization mixture and hybridization at 42°C was allowed to proceed for 12 to 16 hrs. The filters were then washed once sequentially in 2x SSC, 0.1% SDS at 50°C for 20 min, then l x SSC, 0.1% SDS at 55°C for 30 min, and then finally in O.lx SSC, 0.1% SDS at 60°C for 30 min. The filters were exposed to X-ray film (X-OMAT X K - 1 , Kodak). 2.2.8 One dimensional (ID) immunoblot analysis Cells were pelleted by centrifugation and lysed directly in l x Laemmli SDS-PAGE loading buffer (6x Laemmli SDS-PAGE is 350 m M Tris-Cl, pH 6.8, 10% SDS, 600 m M DTT, 0.012% w/v bromophenol blue, 30% glycerol, 12 m M NaF, 12 m M N a 3 V 0 4 , 12 m M EDTA, and complete protease inhibitor cocktail (Roche)). Alternatively, cells in suspension in K K 2 buffer were directly lysed by mixing with 6X Laemmli SDS-PAGE loading buffer in a volume ration of 5:1. A l l samples were boiled at 100°C for 5 min and used for SDS-polyacrylamide gel electrophoresis (Laemmli, 1970) or stored -20°C. Equal loading of proteins was ensured through two methods: (i) determining protein concentrations using a Dc Protein Assay and (ii) through subjecting test samples to SDS-polyacrylamide gel electrophoresis followed by staining the gels with Coomassie blue solution (0.025% w/v Coomassie G-250, 50% methanol, 5% v/v acetic acid) and visual inspection. SDS-polyacrylamide gels consisted of an upper protein stacking gel (3.9% bis-acrylamide (1:19), 125 m M Tris-Cl, pH 6.8, 0.1% SDS, 0.03% ammonium persulphate, 0.1% TEMED) and a lower protein resolving gel (11% bis-acrylamide (1:19), 375 m M Tris-Cl, pH 8.8, 0.1% SDS, 0.03% ammonium persulphate, 0.07% TEMED). A 20 pg protein aliquot of each sample and 2.0 pi of low-range pre-stained SDS-PAGE molecular weight standards (Cat # 161 0305, Bio-Rad) were loaded in each well and electrophoresed at 90V. The electrophoresis buffer was 25 m M Tris, 192 m M glycine, 0.1% SDS, pH 8.3. For electrophoresis prior to the immunoblot analysis of DdCAD-1 phosphorylation, 180 mm x 160 mm x 1 mm size SDS-polyacrylamide gels as above were used. These gels were run at a constant current of 25 mA/gel for 3 hrs with cooling. After electrophoresis, the proteins were transferred to nitrocellulose (Hybond C, Amersham) or PVDF (Hybond P, Amersham) membranes using a Bio-Rad electroblot apparatus (Mini Protean II) submerged in ice-cold transfer buffer (25 m M Tris, 192 m M glycine, 20% 24 methanol) for 1 hr at 90 V (Towbin et al, 1979). The membranes were incubated in 5% powdered milk in Tris-buffered saline (TBS)-Tween (25mM Tris-Cl [pH 8.0], 1.0% NaCl, 0.1% Tween 20) for 1 hr at room temperature and were then washed twice for 5 min with TBS-Tween. Alternatively, all membrane that were to be probed with phospho-specific antibodies were incubated in 3% B S A in TBS-Tween 1 hr at room temperature and were then washed three times for 5 min with TBS-Tween. Membranes were then probed with primary antibodies in 1% w/v powdered milk (Carnation)/TBS-Tween (1/1000 dilutions for phospho-specific antibodies and 1/2000 dilutions for all others) overnight at room temperature or according to the manufacturer's instructions. The membranes were washed three times for 5 min in TBS-Tween and exposed to a secondary antibody (Donkey anti-rabbit or sheep anti-mouse conjugated to horseradish peroxidase, as appropriate) diluted 1/10,000 in TBS-Tween containing 1% powdered milk. Membranes were then washed three times for 5 min with TBS-Tween and bound antibody was detected by an enhanced chemiluminescence assay (ECL, Amersham) using X-ray film (X-O M A T X K - 1 , Kodak). Immunoblots that were directly compared were detected on the same sheet of film to account for differences in exposure time. Where desired, probed membranes were stripped by incubation in 0.2 M NaOH for 10 min, followed by three washes in ddEbO and reblocked as above, before being re-probed with another primary antibody. Blots were scanned using an U M A X - I I scanner ( U M A X , Dallas, Tx, USA) and densitometry was then performed using GeneQuant Analysis software (Molecular Dynamics). 2.2.9 Two dimensional (2D) electrophoresis and immunoblot analysis Cells were lysed in a buffer (Soskic et al, 1999) containing 25 m M Tris-HCl (pH 8.0), 0.3% SDS, 100 m M DTT, 1 m M N a 3 V 0 4 , 1 m M NaF, 2 m M EDTA, 1 m M sodium pyrophosphate, complete protease inhibitor cocktail (Roche), 25 pg/ml DNAse I, and 7 pg/ml RNAse A and centrifuged at 16000 x g for 15 min at 4°C. The supernatant was removed and 80% (v/v) cold acetone added to the supernatant. Proteins were allowed to precipitate on ice for 30 min and then pelleted by centrifugation at 16000 x g for 15 min at 4°C. The protein pellet was dissolved in isoelectric focusing (IEF) sample buffer (8 M urea, 4% (w/v) CHAPS, 1% (v/v) Triton X-100, 1% DTT, 20 m M Tris, 0.5% (v/v) IPG buffer [pH3-10]) and proteins concentrations were determined using a Dc Protein Assay. 25 Ready-to-use Immobiline DryStrips (13 cm, pH 3-10) were re-hydrated overnight in 250 ul of protein solution (750 ug for staining and 250 ug for immunobloting) and IEF was then carried out up to a total of 75 kVh on an IPGphor isoelectric focusing system (Amershan). Prior to SDS gel electrophoresis, focused strips were equilibrated first in SDS equilibration buffer (50 m M Tris-Cl [pH 8.8], 6 M urea, 30% (v/v) glycerol, 2% (w/v) SDS) containing 10 mg/ml DTT for 15 min, and then in the same buffer containing 25 mg/ml iodoacetamide instead of DTT for 15 min. SDS-PAGE was performed on 11% polyacrylamide gels (180 mm x 160 mm x 1 mm) at a constant current of 25 mA/gel for 3 hrs with cooling. Gels were stained with brilliant blue G-colloidal concentrate (Sigma) or silver (Shevchenko et al., 1996). Although some proteins were lost as a result of the acetone treatment, the resolution of components was greatly enhanced by including this step as part of the procedure. For immunoblot analysis, gels were transferred (Towbin et al., 1979) to PDVF membranes, blocked in 3% BSA for 1 hr, and then incubated with either anti-phosphothreonine (1/1000 dilution) or anti-phosphotyrosine (1/1000 dilution) primary antibodies in 3% BSA in TBS-Tween (25mM Tris-Cl [pH 8.0], 1.0% NaCl, 0.1% Tween 20) at 4°C overnight. Membranes were washed in TBS-Tween before being exposed to secondary antibody (donkey anti-rabbit or -mouse IgG conjugated to horseradish peroxidase) diluted 1/5000 in TBS-Tween. Bound antibody was detected by an enhanced chemiluminescence assay (ECL, Amersham) using X-ray film (X-OMAT X K - 1 , Kodak). When comparing levels of phosphoprotein between samples, the chemiluminescence signal on the immunoblots was detected on the same film to prevent differences in exposure time for affecting image analysis. Stained gels and immunoblots were scanned using a UMAX-I I scanner and image analysis/spot matching was performed using Amersham ImageMaster 2D Elite Software or programs available from the proteome 2D-PAGE database web site (http://www.mpiib-berlin.mpg.de/2D-PAGE/). Computer-aided spot matching was performed through the overlaying of the image of an immunoblot, the same immunoblot membrane stained with silver (Sorensen et al., 2002), and a corresponding stained gel. For each phosphoprotein detected on an immunoblot, surrounding landmark proteins were identified from the above overlays and used to locate the position of phosphoproteins on stained gels through the measurement of distances between landmark spot patterns and phosphoprotein spots. A phosphoprotein spot was considered matched i f three independent repeats of this process overlaid the phosphoprotein spot 26 to a single resolved protein on the stained gel. pi values were determined relative to the values specified by the manufacturer of the pi strips (Amersham). 2.2.10 In-gel tryptic protein digestion A l l reagents used for in-gel tryptic digestion (Wilm et al, 1996; Gharahdaghi et al, 1999) were prepared immediately prior to use. Excised protein spots were placed in 1.5 ml siliconized microcentrifuge tubes, which were rinsed with HPLC quality methanol before use. A l l gel pieces were washed in 50% (v/v) methanol and 5% (v/v) acetic acid overnight, after which, the wash solution was removed and the wash repeated for 2 hr. In the case of silver stained spots, gel pieces were destained prior to the wash procedure as described in Gharahdaghi etal (1999). Following the wash procedure, gel spots were digested with trypsin essentially using standard procedures as described in Wilm et al. (1996). Briefly, gel pieces were dehydrated in 200 pi acetonitrile for 5 min and then dried at ambient temperature in vacuum centrifuge for 3 min. 30 pi of 10 m M DTT in 100 m M ammonium bicarbonate was added to the dried gel pieces and incubated for 30 min. After removal of the DTT solution, 30 pi of 100 m M iodoacetamide in 100 m M ammonium bicarbonate was added to the gel pieces and incubated for an additional 30 min. Samples were then dehydrated in 200 pi of acetonitrile for 5 min, then washed in 200 pi of 50 m M ammonium bicarbonate for 10 min, before being dehydrated in acetonitrile and dried. Trypsin was dissolved in ice-cold 50 m M ammonium bicarbonate to a concentration of 25 ng/pl and 30 pi of this solution was then used to re-hydrate the dried gel pieces on ice for 10 mins. Once re-hydrated the excess trypsin solution was removed and the gel pieces were covered with 50 m M ammonium bicarbonate and digested overnight at 37°C. After digestion, peptides were extracted by the addition of 30 pi of 50 m M ammonium bicarbonate and incubation at room temperature for 10 min, followed by centrifugation at 13,000 x g to pellet insoluble material. The supernatant was removed and the procedure repeated two additional times with 30 pi of 50% (v/v) acetonitrile, 5% (v/v) formic acid. The pooled supernatants were reduced to approximately 20 pi by vacuum centrifugation and stored at -80°C 27 2.2.11 Mass spectral identification and analysis Extracted peptides were analyzed by the UVic-Genome B C Proteomics Centre (http://www.proteincentre.com/) on an Applied Biosystems Voyager DE-STR MALDI-TOF and Applied Biosystems/MDS Sciex QStar. Interpretation of MS and MS/MS spectra was done using the program Mascot (Perkins et al, 1999), with MS searches initially done with a maximum mass tolerance of ± 0.2 Da, a single trypsin missed cleavage, and no phosphorylation modification. MS/MS analysis was used to confirm all spot identifications. While MS/MS can be used to detect the phosphorylation of the extracted peptides, this methodology was not used during this analysis. 2.2.12 Microscopy Phase contrast, DIC (differential interference contrast) and epifluorescent images of live amoebae were observed with an Olympus IX-70 inverted microscope equipped with 20x and 40x objective lenses and a 1.5x multiplier. In chemotaxis assays, a Narishige MN-151 joystick style micromanipulator (Narishige Instruments) was used for positioning of micropipet tips. Images were captured with a DAGE-100 CCD camera (Dage Corp) using a Scion LG-3 frame grabber on a Pentium PC running Scion Image 4.0 (Scion Corp). For streaming assays, phase contrast images of live amoebae were observed with an Olympus C K inverted microscope equipped with 4x, lOx and 20x objectives, captured with the DAGE-100 CCD camera (Dage Corp) using a Scion VG-5 frame grabber on a PowerPC Macintosh running Scion Image 1.62 (Scion Corp). For indirect immunofluorescence images, mounted slides were observed on an Zeiss Axioplan fluorescent microscope equipped with 40x, 60x and lOOx objective lenses using a D V C camera (Diagnostic Instruments) and Northern Eclipse imaging software. Images were subsequently complied and edited using Scion Image and Adobe Photoshop 5.0. 2.2.13 Indirect immunofluorescence For indirect immunofluorescence staining for DdCAD-1, cells were deposited on No. l glass coverslips (VWR Scientific Inc.) and allowed to adhere for 10 min in HL5 media. Once adhered, the media was removed and the cells were gently washed with K K 2 to remove the remaining media. Cells were fixed in 4% formaldehyde in K K 2 for 15 min at room temperature. 28 Ax2 cadA~ Figure 4. Immunofluorescent staining of DdCAD-1 in Ax2 and cadA null cells. Ax2 (a, c) and cadA null (b, d) fixed and permeablized with cold methanol before being immunostained by indirect immunofluorescence with anti-DdCAD-1 antibodies (a, b) as described in Materials and Methods. Corresponding DIC images are shown (c, d). Images were taken on an Olympus IX-70 inverted microscope. Bar: 100 um. 29 In the cases where cells were permeablized, the previous solution was removed and cold methanol (-20°C) containing 1% formaldehyde was added for 5 min (Fukui et al, 1987). Nonspecific binding was blocked with 1% (w/v) BSA in PBS for 10 min, before samples were incubated with DdCAD-1 antiserum (1:400 dilution in PBS containing 1% BSA) (Sesaki and Siu, 1996) for 1 hr. Samples were then washed three times in PBS containing 0.05% Tween 20 and then stained with FITC-conjugated goat anti-rabbit IgG (1:200) (Jackson Laboratories) for 1 hr. Samples were then washed three times in PBS containing 0.05% Tween 20 and the coverslips mounted in ProlongR Antifade mounting media (Molecular probes) on Premium microscope slides (Fisher). Images were acquired using a Zeiss Axioplan fluorescent microscope equipped with 60x objective lenses using a D V C camera (Diagnostic Instruments) and Northern Eclipse imaging software. The specificity of DdCAD-1 antiserum (Sesaki and Siu, 1996) was confirmed by staining of cadA null strain {Figure 4). 2.2.14 Cell cohesion assays Cell cohesion assays were performed using a modification (Wong et al., 2002) of the original roller tube assay of Gerisch (Gerisch, 1980). Vegetative cells were centrifuged at 700 x g for 4 min, washed in K K 2 and resuspended in K K 2 at 2 x 107 cells/ml. Cells were starved in rotatory agitated suspension (175 rpm) at 22°C for 4 hrs, after which they were resuspended to a density of approximately 2.5 x 106 cells/ml. Cell aggregates were dispersed by vigorously vortexing for 15 sec. Aggregates were allowed to re-form while rotating on a platform shaker rotating at 180 rpm at room temperature. At indicated times, the number of non-aggregated cells, including singlets and doublets, were scored using a hemocytometer. The percentage of cell aggregation was calculated by subtracting the number of non-aggregating cells (singlets and doublets) from the total number of cells and dividing this number by the total number of cells. The requirement for divalent cations for cell cohesion was assayed in the presence of 10 m M EDTA (Wong et al, 2002). For the visual monitoring of cell cohesion, cells were allowed to adhere together in HL5 media alone or HL5 media supplemented with 10 m M EDTA or 50 pg/ml anti-DdCAD-1 antibodies. After the indicated times, cells were seeded on Nunc tissue culture dishes and images taken on an Olympus C K inverted microscope equipped with equipped with a DAGE-100 CCD camera (Dage Corp). 30 2.2.15 Chemotaxis and aggregation streaming assays Micropipet chemotaxis assays were carried out on vegetative cells that were dispersed at a density of approximately 4 x 105cells/cm2 in HL5 media on Nunc tissue culture dishes. The cells were then rinsed once and submerged in 20% HL5 (Palmieri et al., 2000). At t=0, a micropipet (Eppendorf Femtotip II) filled with 25 m M folate was positioned in the field of view and cell movements monitored by time-lapse microscopy. To observe aggregation streaming, vegetative cells were seeded at approximately 5 x 105 cells/cm2 in Nunc tissue culture dishes and allowed to adhered for 30 min. Cells were washed twice with Bonner's salts (10 NaCl, 10 m M KC1, 2 m M C a C y and then submerged under Bonner's salts. Images were taken on an Olympus C K inverted microscope with the DAGE-100 CCD camera. 2.2.16 Assay of folate stimulation of E R K 2 and PKB phosphorylation For folate stimulation of ERK2/p70 phosphorylation the procedure used by Maeda and Firtel (1997) was utilized. Vegetative cells were washed twice in K K 2 buffer, resuspended at approximately 5 x 106 cells/ml in K K 2 and then agitated at 175 rpm at 22°C for 30 min. For folate stimulations to monitor P K B phosphorylation (Lim, 2002), the above step was excluded. Cells were then washed twice in K K 2 buffer, resuspended in 1 ml of K K 2 at a density of 1 x 10 cells/ml and vortexed gently. Cells were stimulated with folate to a final concentration of 50 nM. At the indicated times, 50 pi samples were lysed by mixing with 10 pi 6x Laemmli SDS-PAGE loading buffer (6x Laemmli SDS-PAGE is 350 m M Tris-Cl, pH 6.8, 10% SDS, 600 m M DTT, 0.012% w/v bromophenol blue, 30% glycerol, 12 m M NaF, 12 m M N a 3 V 0 4 , 12 m M EDTA, and complete protease inhibitor cocktail (Roche)). Protein samples were fractionated by ID electrophoresis and subjected to immunoblot analysis as described in Sections 2.2.8. The antibodies used were the Phospho-p44/42 M A P kinase polyclonal antibody (Cat# 9101) and the phosphothreonine polyclonal antibody (Cat# 9381). Equal loading of proteins was ensured through determining protein concentrations using a D c Protein Assay and checking a duplicate gel with Coomassie G-250. Bound antibody was analyzed using GeneQuant Analysis software (Molecular Dynamics). 2.2.17 Protein translocation assays Cells (1 x 107) were washed in K K 2 buffer and then resuspended in 1 mL K K 2 buffer supplemented with complete protease inhibitors (Roche), 1 m M NaF and 1 m M N a 3 V 0 4 . Cell 31 suspensions were lysed by filtration through a 5.0 pm TMTP membrane and the lysates were collected in microcentrifuge tubes (Diamed). Samples were microfuged at 16,000 x g for 1 min, the sedimented pellet dissolved in 100 pi lx Laemmli SDS-PAGE loading buffer (6x Laemmli SDS-PAGE is 350 m M Tris-Cl, pH 6.8, 10% SDS, 600 m M DTT, 0.012% w/v bromophenol blue, 30% glycerol, 12 m M NaF, 12 m M N a 3 V 0 4 , 12 m M EDTA, and complete protease inhibitor cocktail (Roche)) and subjected to immunoblot analysis as described in Sections 2.2.8. A sample of non-lysed cells was also collected as a 'total protein' control and lysed directly in lx Laemmli SDS-PAGE loading buffer. 2.2.18 Subcellular fractionation For the separation of membranes by differential centrifugation (Bush et al., 1994), 1 x 108 cells were stimulated with folate as described in section 2.2.16 for 8 min. Cells were then pelleted, resuspended in 0.25 M sucrose, 1.0 m M NaF, 1.0 m M N a 3 V 0 4 , 1.0 m M EDTA and broken with a Dounce homogenizer. Under these conditions, 95% of the cells were broken as determined by phase-contrast light microscopy. The homogenate was then subject to differential centrifugation to generation a PI (1000 x g for 5 min) and P2 (10,000 x g for 10 min) membrane fractions. The resulting pellet and supernatant fractions were subjected to SDS-polyacrylamide gel electrophoresis and immunoblot analysis as described in Section 2.2.8. 32 3 R E S U L T S 3.1 The effects of RasG protein levels on discoidin expression 3.1.1 Introduction Preliminary experiments, conducted in collaboration with Meenal Khosla, and described in the Introduction to this thesis, had indicated that activated RasG inhibits discoidin expression. However, the interpretation of the inhibitory effects of RasG(G12T) on discoidin expression was complicated by the fact that the activated RasG protein was itself expressed from the discoidin promoter and therefore could repress its own production. 3.1.2 Accumulation of RasG(G12T) is accompanied by decreased discoidin mRNA levels To independently demonstrate that the presence of RasG(G12T) suppressed expression from the discoidin promoter, transformants were isolated that expressed ra.sG(G12T) under the control of the ribonucleotide reductase (rnrB) promoter (Gaudet and Tsang, 1999). This promoter has been shown to respond to DNA-damaging agents (e.g. methyl methanesulfonate, 4-nitroquinoline 1 -oxide and U V light) by increasing expression, making it useful for the inducible expression of endogenous genes (Gaudet et al., 2001). rnrB-rasG(G\2T) transformants rapidly induced the production of RasG(G12T) protein upon exposure to 10 m M methyl methanesulfonate (MMS) (Figure 5A) and a reduction of discoidin mRNA level was observed after two hrs of M M S treatment (Figure 5B). A similar repression of discoidin mRNA levels was observed at all the cell densities tested (8 x 105 to 4 x 106 cells/ml) (data not shown). When wild-type Ax-2 cells were treated with 10 m M M M S , there was no effect on RasG protein level (Figure 5A) or discoidin mRNA level (Figure 5B). The sensitivity of the discoidin promoter to the presence of RasG(G12T) was further revealed by decreasing the concentration of M M S to 5 mM. While the amount of RasG(G12T) protein that was reduced (Figure 5A), there was still an appreciable repression of discoidin mRNA levels (Figure 5B). These results showed that increased intracellular RasG(G12T) was associated with repressed discoidin mRNA levels. 3.1.3 Discoidin expression in rasG null cells Since discoidin expression was repressed in the presence of elevated levels of RasG(G12T) in the cell, it was anticipated that it would be unaffected, or even enhanced, in rasG 33 (A) Ax-2 rnrB-rasG(G12T) 10 mM MMS 5 mM MMS 10 mM MMS Oh 1h 2h Oh 1h 2h Oh 1h 2h — 34.2 — 28.3 — 19.9 (B) Ax-2 rnrB-rasG(G12T) 10 mM MMS 5 mM MMS 10 mM MMS Oh 1h 2h Oh 1h 2h Oh 1h 2h Figure 5. Effect of R a s G ( G 1 2 T ) expression from the rnrB promoter on discoidin mRNA levels. Ax-2 and rnrB-rasG(G12T) transformed cells were grown in HL5 media to a cell density of approximately 1 x 106 cells/ml and then supplemented with methyl methanesulfonate (MMS) at the indicated concentration. (A) Aliquots were taken either immediately or 1 and 2 hrs after the addition of M M S and RasG protein levels were determined by immunoblot analysis as described under Materials and Methods. Molecular weight markers in kDa are shown. (B) The samples described in (A) were also assayed for discoidin mRNA levels by northern blot analysis. 34 null cells (Tuxworth et al, 1997). In fact, the Discoidin level in rasG null cells was considerably lower than the level in the Ax2 parental line (Figure 6A, lane 7 verses lane 1), suggesting a requirement of RasG for high level discoidin expression. This finding raised the question as to whether the presence of RasG was necessary for any of the previously described aspects of discoidin promoter regulation (Blusch et al., 1992; Burdine and Clarke, 1995; Gomer et al., 1991; Wetterauer et al, 1995). It has been shown that Discoidin levels first increase and then decrease during the growth of Ax2 cells in axenic medium (Wetterauer et al, 1995). The levels of Discoidin in rasG null cells were therefore monitored during axenic growth and compared to the levels in Ax2. The expected pattern of Discoidin expression in Ax2 was observed, with maximum levels of Discoidin occurring during late exponential phase at a cell density of 8.5 x 106 cells/ml (Figure 6A, lanes 1-6). A similar, but less pronounced pattern of discoidin expression was observed in the rasG null cells, with maximum levels of discoidin occurring during late exponential phase for these cells at a cell density of 2.6 x 106 cells/ml (Figure 6A, lanes 7-12). Since more pronounced differences between the two strains might have been masked by a low turnover of the Discoidin protein, discoidin mRNA levels were also monitored. This analysis emphasized the similarities rather than the differences in discoidin expression between the two strains (Figure 6B). Despite the fact that the axenic growth characteristics of the rasG null cells are quite different from those of Ax2 (i.e. rasG null cells growing to lower cells densities in shake suspension) (Tuxworth et al, 1997), the overall expression of discoidin in the two strains in response to axenic growth was remarkably similar. These results suggested that the presence of RasG was not necessary for the axenic growth dependent regulation of discoidin expression, but that RasG did impact discoidin mRNA translation or protein stability (Figure 6A verses 6B). To determine i f folate repressed discoidin expression in the absence of RasG, cells were grown in the presence and absence of folate and Discoidin protein levels determined (Figure 7). Growth in axenic medium supplemented with ImM folate for 24 hrs resulted in a decline in Discoidin protein levels in both control Ax2 cells and rasG null cells, indicative of a repression of discoidin promoter activity in response to folate (Blusch et al, 1992). These results demonstrated that RasG was not necessary for folate repression of discoidin expression. When wild-type Ax2 cells are grown in suspension with bacteria, discoidin expression is suppressed until the bacteria are consumed and the amoebae reach stationary phase (Rathi et 35 (A) Ax-2 rasG null 1 2 3 4 5 6 7 8 9 10 11 12 (B) Ax-2 rasG null 19.9 Figure 6. Discoidin expression in rasG null cells during axenic growth. Ax-2 and rasG null cells were grown in rotatory agitated suspension in HL5 media and aliquots were taken at various times during growth. Under these conditions rasG null cells grew with a generation time of approximately 30 hrs and only reached a cell density of 2 x 106 cells/ml in stationary phase. (A) Discoidin protein levels were determined by immunoblot analysis during growth at the following cell densities (cells/ml): for Ax-2, 1 x 106 (lane 1), 4.2 x 106 (lane 2), 8.5 x 106 (lane 3), 1.2 x 107 (lane 4), 1.8 x 107 (lane 5), 2.2 x 107 (lane 6); for rasG null, 1.2 x 106 (lane 7), 1.4 x 106 (lane 8), 1.9 x 106 (lane 9), 2.3 x 106 (lane 10), 2.6 x 106 (lane 11), 2.8 x 106 (lane 12). Molecular weight markers in kDa are shown. (B) Samples taken at the following cell densities (cells/ml): for Ax-2, 4.2 x 106 (lane 1), 8.5 x 106 (lane 2), 1.2 x 107 (lane 3), 2.2 x 106 (lane 4); for the rasG null, 1.4 x 106 (lane 5) 1.9 x 106 (lane 6), 2.3 x 106 (lane 7), 2.8 x 106 (lane 8) were also assayed for discoidin mRNA levels by northern blot analysis. Equal loading of RNA confirmed by EtBr staining as described in Material and Methods. Figure Credit: Done in collaboration with M . Khosla. 36 Ax-2 rasG null Folate: + + + + Time(hrs) : 0 24 0 24 0 24 0 24 49.5 34.2 28.3 19.9 Figure 7. Folate-repression of discoidin protein levels in Ax2 and rasG null cells. Ax-2 and rasG null cells were grown in HL5 media to a density between 1-2 x 106 cells/ml, washed and then resuspended in fresh HL5 media in the presence (+) or absence (-) of 1 m M folate for 24 hrs. Aliquots were taken at the indicated times and discoidin levels were determined by immunoblot analysis as described in Materials and Methods. Molecular weight markers in kDa are shown. Figure Credit: Done in collaboration with M . Khosla. 37 al.,1991; Rathi and Clarke, 1992). The induction of discoidin expression during this growth period has been attributed to the removal of the inhibitory effects of the bacteria and to increasing levels of PSF in the medium (Burdine and Clarke, 1995). In order to determine i f RasG was necessary for this induction, Discoidin expression in Ax2 and rasG null cells grown in suspension with bacteria was compared (Figure 8). In both strains, Discoidin levels increased when cells had consumed all the bacteria and entered stationary phase (Figure 8), indicating that RasG was not necessary for the inhibition by bacteria and for PSF induction. Discoidin expression is also induced during early development and this induction has been attributed to CMF stimulation (Gomer et al., 1991). To determine i f RasG was necessary for this induction, rasG null cells were washed free of media and plated on agar after growth in suspension with bacteria. The rasG null cells displayed a delay in Discoidin expression relative to that observed for Ax2 (Figure 9). This result was similar to the result described above for the induction of Discoidin during growth on bacteria. The delay in Discoidin expression corresponded to the delay in the aggregation of the rasG null cells (see section 3.3.6), indicating that the presence of RasG was not required for the induction of Discoidin expression normally observed as cells aggregate. Nevertheless, this result indicated that RasG might modulate the induction of Discoidin expression observed during the first six hrs of development. 3.1.4 Summary The expression of activated RasG(G12T) from the rnrB promoter was shown to repress discoidin expression. Despite this repression, RasG was found to be required for high discoidin expression, since discoidin expression was low in rasG null cells. However, RasG was not involved in the regulation of discoidin expression by folate, CMF, PSF, bacteria and growth in axenic media, all signals shown previously to regulate discoidin expression. It should be noted here that although the expression of RasG(G12T) from the discoidin promoter in pVEII-rasG(G\2T) cells has been shown to reduce Discoidin protein levels due to protein turnover (Secko et al., 2001), using the induction times shown in Figure 5, expression of RasG(G12T) from the rnrB promoter did not reduce Discoidin protein levels (D. Secko, unpublished observations). This difference may be due to difference in the time frame of inducible expression, since pVEII-rasG{G\2T) cells are typically induced for 48-72 hrs while rnrB-rasG(G\2T) cells were induced for 2 hrs (Secko et al., 2001). 38 Ax-2 rasG null 4 5 6 7 8 9 10 11 12 49.5 Figure 8. Discoidin expression in rasG null cells during growth on bacteria. Ax-2 and rasG null cells were grown in a suspension of Klebsiella aerogenes as described in Materials and Methods. Cells that had been previously grown in a bacterial suspension for seven-eight generations were inoculated at a density of 5 x 104 cells/ml into a fresh bacterial suspension and discoidin protein levels were monitored during growth at the following cell densities (cells/ml): for Ax-2, 1.6 x 106 (lane 1), 3.5 x 106 (lane 2), 7 x 106 (lane 3), 8.6 x 106 (lane 4), 1.1 x 107 (lane 5), 1.7 x 107 (lane 6); for the rasG null, 1.2 x 106 (lane 7), 3.8 x 106 (lane 8), 5.3 x 106 (lane 9), 7.6 x 106 (lane 10), 8.1 x 106 (lane 11), 8.4 x 106 (lane 12). 39 Ax-2 rasG n u l l 0 3 6 12 0 3 6 12 •49.5 34.2 28.3 Figure 9. Discoidin expression during the development of rasG null cells. Cells were set up to develop on non-nutrient agar buffered with K K 2 after growth in a bacterial suspension to a density of approximately 3 x 106 cells/ml. Protein samples were taken at the indicated times (in hrs) after plating and discoidin protein levels determined. Molecular weight markers in kDa are shown. 40 3.2 Use of an inducible RasG(G12T) expression system to identify Dictyostelium phosphoproteins altered by the expression of RasG(G12T) 3.2.1 Introduction The discovery that RasG regulated discoidin expression revealed an inherent problem with the use of the discoidin promoter to express RasG, and thus the need for an alternative expression system. The studies with the rnrB promoter, outlined in the previous section (3.1), indicated that the effects of RasG(G12T) on discoidin expression were rapid and raised the possibility that this might be a useful system for determining downstream effectors involved in RasG signaling. However, although the rnrB promoter was a useful alternative expression system, it had two shortcomings for additional studies: the need to use D N A damaging agents for induction and the stability of expression. Although the former can be overcome through the use of short induction times {Figure 5), the induction agent does cause cell death over extended exposure times (Gaudet et al., 2001). More troublesome was the observation that RasG(G12T) induction from the rnrB promoter became non-responsive to M M S after the strains had been grown for several passages, ultimately requiring re-transformation and the selection of new transformants (D. Secko, unpublished observations). 3.2.2 Expression of RasG(G12T) from the tetracycline promoter, an alternative expression system Given the constraints of the rnrB promoter system, an alternative Dictyostelium expression system based on the adaptation of the widely used tetracycline (tet) promoter (Blaauw et al., 2000) was tested. To monitor the effectiveness of the tetracycline repressible promoter system the levels of RasG protein in wild-type Ax2 cells containing an empty vector (Ax2::MB) and in cells containing RasG(G12T) under control of the tet promoter (Ax2::MB-ra.s ,G(G12T)) were measured. Levels of discoidin mRNA were also determined. When Ax2::MB and Ax2::MB-ra?G(G12T) cells were grown in the presence of 5 u_/ml tetracycline the expression of RasG protein in these cells was similar (Figure 10A). Comparison of the levels of discoidin mRNA in the Ax2::MB and Ax2::MB-rasG(G12T) cells revealed the discoidin mRNA levels to be slightly repressed Ax2::MB-rasG(G12T) cells (Figure 10A). This suggested that repression of the tet promoter in the presence of 5 Lxg/ml tetracycline was not complete and that potentially 41 (A) Ax2 :MB Ax2::MB-RasG(G12T) hrs: 0 1 2 3 4 0 1 2 3 4 35 — 29 — mm l j t [ L : : i r . . . «* ~» m m - RasG 22 — (B) hrs: 0 1 2 3 4 0 1 2 3 4 . ^ M f e a f e - - i^^Hp I^ ^^ K' discoidin " m R N A Figure 10. Induction of RasG(G12T) protein levels from the tet promoter and its effect on discoidin mRNA levels. (A) Transcription from the tet repressible promoter in Ax2: :MB and Ax2::MB-ra,sG(G12T) cells was induced by washing two times with HL5 to remove tetracycline. Washed cells were resuspended in HL5 and allowed to grow for 4 hrs, during which aliquots were taken at the times indicated and RasG levels determined by immunoblot analysis. Molecular weight markers in kDa are shown. (B) The cell samples taken in (A) were also used for RNA isolation and subsequently assayed for discoidin mRNA levels by Northern blot analysis. 42 undetectable levels of RasG(G12T) were being expressed in the presence of tetracycline. Increases in the levels of RasG were clearly observable in the Ax2::MB-r<zs-G(G12T) cells as early as three hrs after the removal of tetracycline (Figure 10A). This increase was sufficient to induce a repression of discoidin gene expression (Figure 10B). These results revealed that tetracycline repressible promoter system was an effective alternative for the expression of RasG(G12T) and that under these conditions signaling was occurring from RasG to discoidin gene expression. 3.2.3 Detection of protein phosphorylation changes upon the expression of activated RasG The availability of a system for the regulated expression of activated RasG provided an opportunity to further define the molecular components of the RasG signaling pathway. Ras activation in other organisms has been shown to activate phosphorylation cascades, in which proteins are sequentially phosphorylated as signals are transmitted through the cell (Hunter, 2000; Campbell et al., 1998). It was, therefore, hypothesized that the rapid expression of RasG would result in changes in the phosphorylation of proteins that are putative RasG signaling pathway components and/or targets. To determine i f differences in protein phosphorylation could be detected, cells that were either expressing or not expressing activated RasG were induced for three hrs and lysed by the addition of lysis buffer (see Materials and Methods). Ten p.g of protein was fractioned by SDS-P A G E and then transferred to a PDVF membrane. Immunoblot analysis using monoclonal phosphotyrosine or polyclonal phosphothreonine antibodies allowed the detection of a subset of vegetative phosphoproteins (Figure 11). Several proteins exhibited reproducible changes in protein phosphorylation in cells expressing rasG(G\2T) relative to cells containing the empty vector (Figure 11). 3.2.4 Separation of RasG-responsive phosphoproteins by 2D gel electrophoresis To determine the identity of the vegetative proteins exhibiting altered phosphorylation, proteins were first separated by 2D gel electrophoresis, using a pi range of 3-10. These gels were blotted to PVDF membranes and phosphorylated proteins detected once again using monoclonal phosphotyrosine or polyclonal phosphothreonine antibodies (Figure 12). The increased resolution afforded by this method allowed the detection of over 70 phosphothreonine proteins. 43 -111--73--48-33--28 -20-pThr pTyr Figure 11. Effect of RasG(G12T) expression from the tet promoter on the phosphorylation of vegetative proteins. Ax2::MB (lanes 1 and 3) and Ax2::MB-rasG(G12T) (lanes 2 and 4) cells were sampled three hrs after removal of tetracycline and fractionated by ID gel electrophoresis. Immunoblots were assayed for phosphothreonine (lanes 1 and 2) or phosphotyrosine (lanes 3 and 4) containing proteins by using phosphothreonine and phosphotyrosine specific antibodies. Changes in the vegetative phosphorylation pattern caused by RasG(G12T) induction are indicated (closed arrows). 44 Ax2::MB Ax2::MB-RasG(G12T) Figure 12. Immunoblots of vegetative phosphoproteins separated by 2 D gel electrophoresis. Total protein from Ax2::MB (A, C) and Ax2::MB-rasG(G12T) (B, D) cells three hrs after removal of tetracycline was fractionated by 2D electrophoresis and blotted. Membranes were probed with either anti-phosphothreonine (A, C) or anti-phosphotyrosine (B, D) antibodies. Every spot was systematically numbered, irrespective of its detection on other gels and then cross-referenced between gels. The blots shown are representative of three independent experiments and molecular weight (kDa) and pi ranges indicated. For simplicity, only a subset of the detected spots are labeled. 4 5 This is likely to be a minimum estimate of phosphothreonine containing proteins present in vegetative cells, since, for example, s5 and s9 were smeared spots that probably contain several poorly resolved proteins. Consistent with this interpretation, it has been observed previously that high molecular mass components are difficult to resolve (Gorg et al., 2000). Only three phosphotyrosine containing proteins were detected (Figure 12), suggesting that the method was not sufficiently sensitive to detect most species. The predominant phosphotyrosine protein detected in ID SDS-PAGE gels (Figure 11) was resolved into two spots, sl37 and sl38 (Figure 12), both of which showed an increase in tyrosine phosphorylation in the Ax2::MB-rasG(G12T) cells. The phosphoproteins could be subdivided into three categories: those exhibiting unchanged intensity, those exhibiting altered intensity, and those exhibiting apparent pi shifts, as assessed by alterations in position, and examples of each category are shown in Figure 13. The majority of phosphoproteins detected fell into the first category, those not showing any significant change in response to RasG(G12T) induction, e.g. s60 and s61 (Figure 13A). Of the proteins in the second category, some showed a detectable level of phosphorylation in uninduced cells and were then further phosphorylated or dephosphorylated in response to activated RasG; for example, proteins s20 and s35 were present in uninduced cells, but exhibited a marked reduction in phosphorylation in response to RasG(G12T) expression (Figure 13B). For some proteins in this category, phosphorylation increased from or decreased to undetectable levels in response to the activation of RasG. For example, protein s98 was not detectably phosphorylated in uninduced cells but was significantly phosphorylated in response to activated RasG (Figure 13C). A few proteins fell into the third category as being phosphorylated in both strains, but also exhibiting an apparent shift in pi in response to activated RasG, e.g. sl38 (Figure 13D), although identification of both spots will be essential to confirm that a pi shift has occurred. It should be noted that sl38 was also a category 2 protein, since the spot increased appreciably in intensity in the induced cells. Nineteen phosphoproteins (3 phosphotyrosine and 16 phosphothreonine), shown in Figure 12, displayed an appreciable and reproducible alteration in phosphorylation level and were considered for further study. Some of the changes in phosphorylation were very large (e.g. the increased phosphorylation of s!37), while some were more modest (e.g. the increased 46 (A) (B) (C) (D) Ax2::MB 6.7—pi—6. 33 MW 26 44 I MW 42 5.5—pi—5.8 5.1—pi —5.4 Ax2::MB-RasG(G12T) 6.7—pi—6.8 Figure 13. Representative changes in phosphorylation between the Ax2::MB and Ax2::MB-rasG(G\TY) strains. Selected phosphoproteins are shown as image expanded blots. Molecular weight (kDa) and pi ranges are shown. (A) Proteins whose phosphorylation was unchanged by RasG(G12T) induction. (B and C) Proteins whose phosphorylation was altered (D) A phosphoprotein showing a possible shift in pi. 47 100 80 60 40 20 H a. i J i 5 136 33 19 55 53 25 35 8 21 20 56 102 36 78 18 Spot Number 57 138 137 Figure 14. Quantification of protein phosphorylation levels between the Ax2::MB and Ax2::rasG'(G12T) strains. 2D immunostained spots detected with anti-phosphothreonine or anti-phosphotyrosine antibodies (Figure 10) were quantified, their intensities normalized to an unchanged 2D reference spot on each gel. In addition, standard markers on each gel were used to check this normalization. The relative intensity of a subset of spots is indicated. Values plotted are the means and standard deviations for three experiments. Only proteins that showed a reproducible alteration in the three independent experiments are shown. Ax2: :MB, white bars; Ax2::MB-rasG(G12T), gray bars. 48 phosphorylation of sl8 and the decreased phosphorylation of sl9) (Figure 14). Since these proteins represented only those most easily detected, there is clearly a complex pattern of protein phosphorylation or dephosphorylation in response to activated RasG. One component that did not exhibit an appreciable change (s57) was included in the analysis, since it appeared to be a major phosphorylated species. 3.2.5 Identification of putative RasG-responsive phosphoproteins Of the nineteen phosphoproteins displaying changes in phosphorylation, only thirteen were clearly resolved by 2D electrophoresis into individual proteins and these were chosen for identification by mass spectrometry. The degree of resolution of these proteins was determined by matching the immunoblots to a corresponding brilliant blue G-colloidal or silver stained gel (Figure 15) as described under Materials and Methods. This allowed an estimation of the amount of a protein of interest and provided a measure of the reproducibility of the separation of neighboring proteins. Although the silver stained immunoblots contained a lower spot density than the stained gel, their overlay with the original immunoblot proved useful for determining the location of the proteins. For each of these thirteen proteins, the corresponding brilliant blue G-colloidal or silver stained spots did not noticeably change in staining intensity upon induction of activated RasG (Figure 15), indicating that the changes in phosphorylation were not due to differences in protein levels. As was found with rnrB-rasG(G\2T) cells (Section 3.1.4), expression of RasG(G12T) from the tet promoter under the time frame utilized did not affect Discoidin protein levels. These 13 proteins were excised from brilliant blue G-colloidal or silver stained gels, in-gel digested with trypsin (Wilm et al., 1996) and the peptides extracted from these digests were used for peptide mass fingerprint analysis. Proteins present in low abundance, as revealed by weak silver stain intensity, were pooled from up to five individual gels to obtain adequate material for identification. Mass spectral identifications were accomplished by both peptide mass fingerprinting and MS/MS analysis (Graves and Haystead, 2002). Matching the obtained mass data against the Dictyostelium database (http://dictybase.org/index.html) allowed an identification of all 13 spots (Table 1). When selected peptides from each sample were subsequently subjected to MS/MS analysis to generate peptide sequence data, the results 49 A x 2 : : M B A x 2 : : M B - R a s G ( G 1 2 T ) 3 pi 10 3 pi 10 Figure 15. Brilliant blue G-colloidal stained 2D gels of vegetative Ax2 : :MB and A x 2 : : M B -ras(j(G12T) lysates. A representative example of the 2D separation of acetone-precipitated vegetative cell lysates is shown for the strains Ax2: :MB and Ax2::MB-ra5 ,G(G12T). Molecular weight markers (kDa) and pi ranges are indicated. 50 Table 1: A list of Ras-responsive phosphoproteins identified in this study Known/ Spot Identified Predicted Number Protein Function Mass pi % Genbank (Da) Sequence3 Accession Number Signaling Proteins 8 RasGEF-R 20 Protein Kinase B 102 Phosphatidylinositol transfer protein 1 21 Putative cell division protein FtsZA 70959 51315 31365 7.23 6.42 5.84 56854 5.42 28 22 23 29 AAN46870 AAA76692 AAF74409 AAG37880 Adhesion/ Cytoskeletal 55 Calcium-dependent cell 24139 5.40 adhesion moleclue-1 56 Calcium-dependent cell adhesion moleclue-1 24139 5.40 57 Calcium-dependent cell 24139 5.40 adhesion moleclue-1 137 Actin 41803 5.23 138 Actin 41803 5.23 73 55 37 30 P54657 P54657 P54657 A T D O A T D O Translation/ Metabolism 19 98 Similar to seryl-tRNA 52058 6.13 synthetase, cytoplasmic 192908 5.85 Putative UDP-glucose:glycoprotein glucosyltransferase precursor Pyruvate dehydrogenase 39217 5.69 E l component beta subunit 46 20 A C 117072 AAM08766 40 AAO52409 Unknown 78 Hypothetical protein 67328 9.58 14 AAO52604 a % sequence equals the percetage of a protein sequence matched by peptide mass fingerprinting (Graves and Haystead, 2002) confirmed the database identification. In two cases, the same protein was identified in more than one spot, e.g. s55, s56 and s57 (Calcium-dependent cell adhesion moleclue-1 or DdCAD-1), and sl37 and sl38 (Actin), suggesting that these proteins were phosphorylated on more than one site. Therefore, the thirteen identified spots totaled ten different proteins. Of the phosphoproteins identified as being RasG-responsive, several were signaling molecules, including P K B and a RasGEF, proteins that have been shown to be involved in Ras signaling pathways in other cells, as well as Phosphatidylinositol Transfer Protein 1 (PITP1) and FtsZA. The remaining proteins were: a cell adhesion protein (DdCAD-1) and the cytoskeletal protein, actin; two proteins involved in metabolism (a pyruvate dehydrogenase and UDP-glucose:glycoprotein glucosyltransferase); a seryl-tRNA synthetase, and a protein of unknown function. Five of the proteins showed an increase in phosphorylation in response to RasG(G12T) expression; Actin [sl37, sl38], Pyruvate Dehydrogenase [s98], Seryl-tRNA Synthetase [sl8], PITP1 [sl02], and the unknown protein [s78]. Four proteins showed a decrease in phosphorylation in response to RasG(G12T) expression; RasGEF-R [s8], P K B [s20], FtsZA [s21], and UDP-glucose:glycoprotein glucosyltransferase [sl9]. One protein, D d C A D l showed a complex pattern of phosphorylation changes, indicating multiply phosphorylated forms of the protein. These forms exhibited either an appreciable decrease (s55 and s56) or a slight increase (s57) in phosphorylation response to RasG(G12T) expression. These alterations in phosphorylation did not correlate with predicted function, as both increases and decreases in phosphorylation appeared in each functional category (compare Table 1 with Figure 14). Furthermore, these proteins are not expected to all be components of a single RasG pathway, since RasG has been suggested to regulate multiple signaling pathways (Zhang et al., 1999) and mammalian Ras proteins interact with multiple effectors to regulate several signaling pathways (Shields et al., 2000). 3.2.6 Total protein level of a subset of RasG-responsive phosphoproteins does not change upon RasG(G12T) expression The detection of phosphorylation differences in the identified RasG-responsive phosphoproteins (Table 1) could be interpreted in two ways: (i) as changes in activity of kinases or phosphatases in the cell that result in altered phosphorylation or (ii) changes in the total 52 protein level of these targets. As mentioned in the previous section, the brilliant blue G-colloidal or silver staining intensity between the Ax2: :MB and Ax2::MB-ra,sG(G12T) strains did not significantly change upon induction {Figure 15), suggesting that the alterations in phosphorylation observed were not the result of changes in protein levels. To further confirm this, Ax2: :MB and Ax2::MB-rasG(G12T) cells were sampled after three hrs of induction and lysed directly in SDS-PAGE lysis buffer. Ten pg of protein were then loaded onto a ID SDS-P A G E gel, subjected to electrophoresis and transferred to a PDVF membrane. Immunoblot analysis was carried out to detect the total protein levels of three of the RasG-responsive phosphoproteins identified in Table 1, namely Actin, P K B , and DdCAD-1 (Figure 16). As expected, the levels of RasG protein increased in the Ax2::MB-ra,sG(G12T) cells upon removal of tetracycline, but in contrast, the levels of Actin, P K B and DdCAD-1 all remained constant. Additionally, the level of RasC, a protein that was not expected to be affected by RasG(G12T) induction, provided an additional control (Figure 16). These results confirmed that for a subset of the identified RasG-responsive proteins the levels of total protein did not change, further suggesting that the observed changes were due to differences in phosphorylation. 3.2.7 Confirmation of the identities of RasGEF-R and DdCAD-1 The gene encoding RasGEF-R was initially identified by the Insall laboratory as a cDNA clone, SSD492, whose predicted product showed strong homology to the Dictyostelium RasGEF-A (Aimless) and to other putative RasGEF proteins (Wilkins et al., 2001). The Insall laboratory subsequently went on to create three independent isolates that were null for RasGEF-R protein. The availability of these strains allowed confirmation that RasGEF-R was indeed one of the phosphothreonine proteins. Total protein samples taken from the gefR null strain, along with a random integrant control (IR26), were subjected to 2D immunoblot analysis. The s8 immuno-spot identified as RasGEF-R was found in the random integrant strain as before, but was not observed in the gefR null strain (Figure 17). The corresponding silver stained protein spot that had been used for mass spectrometric identification of s8 was also absent (Figure 17). Together these results support the conclusion that s8 is in fact RasGEF-R (Secko et al., 2004). A strain null for DdCAD-1 has also been created (Wong et al, 2002) and the availability of this strain allowed a similar spot identity confirmation. Total protein samples taken from the 53 1 2 4 9 -2 9 -49 2 9 -- Actin - DdCAD-1 - P K B - R a s G 29 - R a s C Figure 16. Total protein levels of three putative RasG-responsive phosphoproteins in a RasG(G12T) expressing and vector control strain. Total protein from Ax2::MB (1) and Ax2::MB-nzsG(G12T) (2) cells three hrs after removal of tetracycline was fractionated by ID electrophoresis. Total protein levels of Actin, DdCAD-1, PKB, RasG and RasC were detected by immunoblotting with specific antibodies to each protein as described in Materials and Methods. Molecular weight markers (kDa) are indicated. 54 IR26 fifefR" pThr Immunoblot silver-stained 2D gel Figure 17. Examination of the gefR' and parental control strains for the presence of RasGEF-R phosphoprotein. Total protein from a gefR and parental control strain (IR26) was fractionated by 2D electrophoresis. Blotted membranes were probed with anti-phosphothreonine antibody to detect phosphoproteins. The section of the 2D immunoblot and a corresponding silver-stained 2D gel known to contain the s8 immunospot, identified as RasGEF-R, is shown along with two additional spots (s6, s7) for reference. Molecular weight (kDa) markers and pi ranges are indicated. 55 Ax2 cadA' mm / i \ 55 56 57 5.5 — pi—5.8 5.5 30 pThr immunoblot silver-stained 2D gel Figure 18. Examination of the cadA' and Ax2 strains for the presence of DdCAD-1 phosphoprotein. Total protein from cadA' and Ax2 strains was fractionated by 2D electrophoresis and blotted membranes were probed with anti-phosphothreonine antibody to detect phosphoproteins. The section of the 2D immunoblots and silver-stained 2D gels known to contain the s55, s56, and s57 immunospots, identified as DdCAD-1, are shown. Additional silver-stained spots (a, b, c, d) are labeled for reference. Molecular weight (kDa) markers and pi ranges are indicated. 56 cadA null strain (cadA'), along with a control Ax2 strain, were subjected to 2D immunoblot analysis. The s55, s56, and s57 immuno-spots identified as DdCAD-1 were all found in the Ax2 strain as before, but not observed in the cadA null strain (Figure 18). The corresponding silver stained protein spots that had been used for mass spectrometric identification of s55, s56, and s57 were also absent (Figure 18). Together these results suggest that s55, s56, and s57 are in fact DdCAD-1. For clarity these phosphoproteins have therefore been re-named DdCAD- la (s57), DdCAD-lb (s56) and DdCAD- lc (s55). 3.2.8 Summary Rapid RasG(G12T) expression from the inducible tetracycline promoter was found to alter the phosphorylation of a sub-set of vegetative phosphoproteins. Thirteen of these phosphoproteins were recovered from 2D gels, in-gel digested with trypsin, and identified by mass spectrometry. Since some of these components represented the same protein in different phosphorylation states a total of ten putative RasG-regulated signaling targets were identified by this analysis. Total protein levels for a subset of these identified proteins were found not to change upon Ras(G12T) induction, indicating that the observed phosphorylation changes were not due to changes in protein synthesis of protein degradation, and suggesting the existence of as-yet-unidentified RasG regulated kinases and/or phosphatases during vegetative growth. Two of these phosphoproteins, RasGEF-R and DdCAD-1, were shown to be absent in their respective null strains. Although, it cannot be ruled out that the loss of RasGEF-R and DdCAD-1 affects the expression or phosphorylation of these phosphoproteins, resulting in their absence in each respective null strain, the data tentatively identified the spots s55, s56, and s57, as DdCAD-1 and the spot s8 as RasGEF-R. 57 3.3 Activated RasG and cell-to-cell adhesion in Dictyostelium 3.3.1 Introduction The identification of DdCAD-1, a known developmental adhesion molecule in Dictyostelium (Coates et al., 2001), as a RasG-responsive phosphoprotein (Figure 18) raised the question as to whether this phosphorylation in any way affects cell adhesion. 3.3.2 Expression of RasG(G12T) during vegetative growth is accompanied by cell clumping Vegetative Dictyostelium cells are normally not cohesive when grown in shake suspension on HL5 media, but upon starvation and the initiation of development cells start to adhere to each other and DdCAD-1 is involved in this initial adhesion (Coates et al, 2001). When Ax2::MB and Ax2::MB-rasG(G12T) cells were grown in shake suspension in the presence of 5 pg/ml tetracycline they grew as individual cells and were not cohesive (Figure 19). In contrast, when Ax2::MB-rasG(G12T) cells were grown in the absence of tetracycline to allow the induction of RasG(G12T) expression, some cells formed clumps, whereas, Ax2: :MB cells grown in the absence of tetracycline did not clump (Figure 19), indicating that the induction of cell-to-cell adhesion was due to the increased levels of RasG(G12T) and not the removal of tetracycline. DdCAD-1 is expressed during vegetative growth in axenic media (Figure 16; Coates et al., 2001), although it is not normally involved in mediating cell-to-cell adhesion until the on-set of early development (Sesaki and Sui, 1996). The finding that RasG(G12T) expression caused cell cohesion during growth raised the possibility that the activation of RasG signaling was prematurely inducing cell adhesion through the dephosphorylation of DdCAD-1. 3.3.3 RasG(G12T)-induced vegetative cell clumping can be blocked with DdCAD-1 inhibitors To test whether the RasG(G12T)-induced cell clumping was mediated by DdCAD-1 two properties of DdCAD-1 -mediated adhesion were examined. First, DdCAD-1 adhesion requires Ca and is thus sensitive to calcium chelators like EDTA (Wong et al., 1996) and second, antibodies specific to DdCAD-1 have been shown to block cell-to-cell adhesion 58 Ax2::MB Ax2::MB-RasG(G12T) + Tet ° . <c_______ Tet Figure 19. Induction of cell-to-cell adhesion during vegetative growth in a RasG(G12T) expressing strain. A x 2 : : M B and Ax2::MB-ra.sG(G12T) cells, grown in H L 5 media in the presence of 5 ug/ml tetracycline (+ Tet) or absence of tetracycline (-Tet) for three hrs, were photographed on an Olympus C K inverted microscope. The arrow indicates an example of the cell-to-cell adhesion observed in Ax2::MB-rasG(G12T) cell cultures washed free of tetracycline. Bar: 100 um. 59 mediated through this molecule (Brar et al., 1993). Ax2::MB and Ax2::MB-rasG(G12T) cells were grown in the absence of tetracycline for three hrs to allow the induction of RasG(G12T) in the presence of either 10 m M EDTA or 50 pg/ml anti-DdCAD-1 antibodies. Control cells incubated in the absence of EDTA or anti-DdCAD-1 antibodies formed clumps as before (Figure 20). The addition of either EDTA or antibodies specific for DdCAD-1 to the media prevented Ax2::MB-rasG(G12T) cells from adhering (Figure 20), consistent with the idea that the observed vegetative cell-to-cell adhesion induced by RasG(G12T) expression is mediated by DdCAD-1. 3.3.4 DdCAD-1 localizes to the cell surface upon the expression of RasG(G12T) During vegetative growth in axenic media, DdCAD-1 is localized in the cytoplasm and the absence of DdCAD-1 at the cell surface may explain why vegetative cells are not adhesive (Sesaki and Sui, 1996; Sesaki et al., 1997). However, during development, as cells become more cohesive, some DdCAD-1 becomes localized at the cell surface (Sesaki and Sui, 1996; Sesaki et al, 1997). The observation that cells expressing RasG(G12T) tend to be adhesive during vegetative growth raised the possibility that DdCAD-1 was re-localized to the surface upon the expression of activated RasG. Indirect immunofluorescence microscopy using anti-DdCAD-1 antibodies (Sesaki and Siu, 1996) was undertaken to determine the localization of DdCAD-1 in induced Ax2: :MB and Ax2::MB-ra^G(G12T) vegetative cells (Figure 21). In vegetative Ax2::MB cells that had been permeablized with cold methanol (Fukui et al, 1987) a diffuse cytoplasmic localization of DdCAD-1 was observed (Figure 21a), that was similar to the previously reported result (Sesaki and Sui, 1996). In contrast, permeablized vegetative Ax2::MB-rasG(G12T) cells additionally contained intensely stained rings of DdCAD-1, that were clearly observable in individual cells (Figure 21b), although somewhat masked in cell clumps due to the increased background staining (Figure 21c). To restrict the staining to DdCAD-1 localized at the cell surface, non-permeablized cells were subjected to indirect immunofluorescence microscopy with the anti-DdCAD-1 antibodies. Under these conditions, vegetative Ax2::MB cells were largely unstained (Figure 21d), consistent with DdCAD-1 being localized predominately in the cytoplasm in these cells. In contrast the DdCAD-1 ring structures were once again observed in the vegetative Ax2: :MB-60 (A) f (B) (C) Ax2::MB Ax2::MB-RasG(G12T) Control 1mM EDTA anti-DdCAD-1 antibodies Figure 20. Disruption of RasG(G12T) mediated cell-to-cell adhesion with inhibitors of DdCAD-1 adhesion. Ax2::MB and Ax2::MB-rasG(G12T) cells were washed and incubated for three hrs in tetracycline-free media with either no additions (A), 1 mM EDTA (B), or 50 pg/ml anti-DdCAD-1 antibodies (C). Cells were photographed on an Olympus C K inverted microscope. Bar: 100 pm. 61 Figure 21. Immunofluorescent localization of DdCAD-1. Ax2::MB (a, d) and Ax2: :MB-ra^G(G12T) (b, c, e) cells were grown in the absence of tetracycline for three hrs. Cells were then fixed and either permeablized with cold methanol (a, b, c) or not permeablized (d, e), before being immunostained by indirect immunofluorescence with anti-DdCAD-1 antibodies as described in Materials and Methods. Images were taken on a Zeiss Axioplan Fluorescent Microscope. Bar: 5 pm. 6 2 rasG(G\2T) cells (Figure 21e), strongly suggesting that DdCAD-1 was localized to the cell surface of these cells. This observation revealed that the activation of RasG resulted in the re-localization of DdCAD-1 and offers an explanation as to why the vegetative Ax2: :MB-rasG(G12T) cells tended to be more cell-to-cell adhesive than the control cells. 3.3.5 Cell adhesion during early developmental is enhanced in RasG(G12T) expressing cells DdCAD-1 mediated adhesion has been proposed as the major adhesion process present soon after the on-set of starvation (Coates et al., 2001), with other adhesion systems (e.g. gp80 and gpl50) induced several hrs later (Desbarats et al, 1994). Gerisch's original analysis of this process made use of a rolling tube cohesion assay to show the ability of early developing Dictyostelium cells to adhere to one another (Gerisch, 1968). This cell cohesion assay was later adapted by Sui and colleagues to assess the ability of strains to adhere via DdCAD-1 contacts sites during early development (Lam et al., 1981; Wong et al., 2002). This standard cell cohesion assay was preformed on Ax2::MB-rasG(G12T) and Ax2::MB cells that had been washed free of tetracycline and starved for 4 hrs (Figure 22), a time when only EDTA-sensitive DdCAD-1 dependent cell adhesion is occurring (Wong et al., 2002). The cells were dissociated by vigorous vortexing so that cells were not in aggregates at time zero and then allowed to re-associate. Some re-association of the Ax2::MB cells was detected 20 minutes after their dissociation (Figure 22). However, the re-association of induced Ax2: :MB-rasG(G12T) cells was much more rapid, with significant adhesion occurring within 5 minutes after the cells had been vortexed (Figure 22). The addition of 10 m M EDTA during the assay completely prevented re-association of both sets of cells, indicating that the observed re-association was in fact due to EDTA-sensitive adhesion. These results indicated that cells expressing RasG(G12T) were not only more cohesive during vegetative growth but also exhibited an enhanced ability to adhere to each other during early development. 3.3.6 DdCAD-1 phosphorylation decreases during Dictyostelium development Since there was a decrease in DdCAD-1 phosphorylation in cells expressing activated RasG, the forgoing results suggested that there was an inverse relationship between DdCAD-1 phosphorylation and DdCAD-1 mediated cell adhesion. Since wild-type cells became more 63 Ax2::MB-rasG(G12T) Ax2::MB Ax2::MB-rasG(G12T) + EDTA Ax2::MB + EDTA Time (mins) Figure 22. Cell-to-cell cohesion in developing Ax2::MB and Ax2::MB-rasG(G12T) cells. A x 2 : : M B and Ax2::MB-ra.sG(G12T) cells were washed free of tetracycline and starved for four hrs in K K 2 in shake suspension. Cells were subsequently disassociated by vortexing and cell re-association monitored over time with or without the addition of 10 m M EDTA. Cells are not in aggregates at time zero and the percentage of cell re-association was calculated by scoring non-aggregated cells with a hemocytometer (% Aggregation = [total number of cells - non-aggregating cells] / total number of cells). Data represents mean of three independent experiments ± standard deviations. (Ax2: :MB [•], A x 2 : : M B + EDTA [•], A x 2 : : M B -rasG(G12T) [•], Ax2::MB-rasG(G12T) + E D T A [A]). 64 adhesive through DdCAD-1 contact sites as development proceeds (Sesaki and Sui, 1996; Sesaki et al., 1997), it was important to determine whether DdCAD-1 phosphorylation decreased during development. Because of the technical difficulties involved in processing multiple samples for comparison by 2D electrophoresis, this analysis was performed using ID electrophoresis. Ax2: :MB cells washed free of tetracycline and starved had relatively constant levels of phosphorylated DdCAD-1 during the first 4 hrs of development, after which the levels of phosphorylated DdCAD-1 decreased (Figure 23). In contrast, Ax2::MB-ra.sG(G12T) cells had a level of phosphorylated DdCAD-1 throughout the starvation period that was similar to that for the Ax2::MB cells that had been starved for 8 hrs (Figure 23). Both sets of cells had similar levels of total DdCAD-1 throughout the starvation period (Figure 23), indicating that the observed phosphorylation changes were not due to changes in protein level. cadA null cells were included to control for other phosphothreonine proteins that might co-migrate with DdCAD-1 during the ID electrophoretic procedure. In the cadA null samples a low intensity band that increased during development migrated in the same position as DdCAD-1 (Figure 23). The gels shown in Figure 23A were therefore quantitated by densitometry, so that the contribution of the non-DdCAD-1 phosphorylation to the phosphoprotein signal could be subtracted. This analysis suggested that the decrease in DdCAD-1 phosphorylation for the Ax2: :MB was somewhat greater than the uncorrected data showed (Figure 23B) and that a slight decrease in DdCAD-1 phosphorylation in the Ax2::MB-ra.s'G(G12T) cells may have occurred during the course of the 8 hr starvation (Figure 23B). Although not quantited by densitometry, a similar decreasing trend in DdCAD-1 phosphorylation was also observed for an independent repeat of this experiment. 3.3.7 A correlation between the decrease in DdCAD-1 phosphorylation and the cellular localization of DdCAD-1 DdCAD-1 is known to move from the cytoplasm to the cell surface as development proceeds (Sesaki and Sui, 1996) and it was of interest to determine in this movement correlated with the observed decrease in DdCAD-1 phosphorylation. To investigate this Ax2: :MB and Ax2::MB-nzsG(G12T) cells were stained for DdCAD-1 at 8 hrs after the onset of starvation (Figure 24). As Sesaki and Sui (1996) had reported, DdCAD-1 was diffusely localized to the cytoplasm at the onset of development in Ax2: :MB cells, and after 8 hrs of starvation was more localized to the cell periphery, which correlated with the large decrease in DdCAD-1 65 (A) Ax2::MB Ax2::MB-RasG(G12T) cad A' hrs: 0 2 4 6 8 29 — 0 2 4 6 8 0 2 4 6 8 29 — (B) pThr immunoblot -a DdCad-1 immunoblot Time (hours) Figure 23. Levels of DdCAD-1 phosphorylation during development. Ax2: :MB, Ax2::MB-rasG(G12T) and cadA' cells were starved in K K 2 in shake suspension in the absence of tetracycline. (A) At the indicated times, protein samples were fractionated by ID electrophoresis and immunoblots probed with anti-phosphothreonine antibodies to detect phosphorylated DdCAD-1 (closed arrow). Blots were then stripped of bound antibody and re-probed with anti-DdCAD-1 antibodies to detect total DdCAD-1 levels (open arrow). Molecular weight (kDa) markers are indicated. (B) The immunoblots probed with anti-phosphothreonine antibodies shown in (A) were quantified by densitometric analysis. The values observed for cadA' cells were subtracted from the phosphorylated DdCAD-1 for Ax2: :MB (•) and Ax2::MB-rasG(G12T) (•) cells and normalized relative to amount of phosphorylated DdCAD-1 in Ax2::MB cells at 0 hrs. 66 Ax2::MB Ax2::MB-RasG(G12T) Figure 24. Comparison of DdCAD-1 localization during development in Ax2::MB and Ax2::MB-rasG(G12T) cells. Ax2::MB (a, c) and Ax2::MB-ra.sG(G12T) (b, d) cells were washed free of tetracycline and starved in K K 2 in shake suspension for the indicated times. They were then fixed and immunostained by indirect immunofluorescence with anti-DdCAD-1 antibodies as described in Materials and Methods. Images were taken on a Zeiss Axioplan Fluorescent Microscope. Bar: 5 um. 67 phosphorylation (Figure 21). In Ax2::MB-ras ,G(G12T) cells expressing RasG(G12T), the DdCAD-1 rings structures present at the onset of starvation became replaced with more general membrane staining after 8 hrs of starvation (Figure 24), also correlating with the smaller decrease in DdCAD-1 phosphorylation (Figure 23). 3.3.8 rasG null cells are defective in streaming during aggregation DdCAD-1 has been implicated in mediating side-to-side contact between cells during aggregation (Coates et al, 2001) and the above findings suggest that RasG may partly regulate this function. Since RasG(G12T) expressing cells do not develop (Khosla et al., 1996) and therefore do not form aggregation streams in which side-to-side contacts can be monitored, the effects of RasG(G12T) on aggregation stream formation could not be monitored. However, rasG null cells do develop (Tuxworth et al., 1997) and I was therefore able to assess aggregation stream formation by these cells. Since cells expressing RasG(G12T) show increased cohesion (see Section 3.3.5), it was hypothesized that rasG null cells would show defective side-to-side contacts indicative of a reciprocal decrease in cell cohesion during early development. Cellular streaming was observed in cell monolayers washed free of nutrient media and submerged under a non-nutrient solution (Bonner's salts). Ax2 cells began to form aggregation streams after 8 hrs under these conditions and by 12 hrs had begun to form aggregation centers (Figure 25). rasG nulls cells did not begin to form aggregation streams until 12 hrs (Figure 25). This result was consistent with the approximately 4 hr delay observed in the initiation of developmental Discoidin expression discussed previously (Figure 9). At 14 hrs, rasG null cells had begun to form aggregation centers, but these were more disorganized than in the wild type and multiple aggregation centers were evident (Figure 25A, bottom panel, 14hr). For comparison, cadA null cells were included in this analysis (Figure 25A, middle panel). cadA null cells also formed cellular streams that were less organized than those of the wild type and multiple small aggregation centers, a phenotype similar to that for the rasG null cells. In addition, the aggregation of cadA null cells was delayed by approximately 2 hours (Figure 25A, middle panel). These results indicated that rasG null cells are defective in developmental streaming, providing further evidence that RasG might be important in the regulation of side-to-side contacts during aggregation (Sesaki and Siu, 1996) by regulating DdCAD-1 phosphorylation. In 68 (A) Ax2 cadA rasG Ohr 8hr Ohr "J WEE**-: - : 8hr 8hr Figure 25. 69 (B) Ax2 rasG Figure 25. Cel lu lar streaming in Ax2, rasG nu l l and cadA nu l l cells. Cells were seeded as monolayers at a density of 5 x 105 cells/cm5 in Nunc tissue culture dishes and submerged under Bonner's salts solution. (A) Images were photographed at the indicated time (hrs) after the initiation of starvation. Bar: 100 pm. (B) Aggregation streams corresponding to 8 hrs for Ax2 and 12 hrs for the rasG null were also photographed at higher magnification. Cells were photographed on an Olympus C K inverted microscope. Bar: 50 pm. 70 addition, these results revealed a role for RasG in regulating developmental timing during early development. 3.3.9 RasG is required for high levels of DdCAD-1 expression Since rasG nulls cells were defective in developmental streaming, a process known to involve DdCAD-1 mediated adhesion, and RasG(G12T) expressing cells exhibited increased developmental adhesion and decreased DdCAD-1 phosphorylation, it was anticipated that the DdCAD-1 phosphorylation would be increased in rasG null cells. The level of DdCAD-1 phosphorylation was therefore measured in vegetative rasG null cells as compared to a vegetative Ax2 control strain (Figure 26). Total protein samples, taken from the rasG null strain (rasG~) and the control Ax2 strain, were subjected to 2D immunoblot analysis. A l l three DdCAD-1 phosphoproteins were found in the Ax2 strain as before, but surprisingly there was greatly reduced phosphorylation of all three DdCAD-1 species in the rasG null strain (Figure 26A). However, the total amount of DdCAD-1 was greatly decreased in rasG null cells in comparison to Ax2 control cells (Figure 26B), despite the fact that Discoidin protein levels do not change in response to the short term activation of RasG. These results suggest that the decrease in DdCAD-1 phosphorylation was due to decreased DdCAD-1 expression levels and indicate that RasG is an important regulator of DdCAD-1 expression during vegetative growth in axenic media. Furthermore, the results suggested that the similarity between the streaming defects in the rasG and cadA null strains could have been due to the reduction in the DdCAD-1 level in rasG null cells, in addition to an alteration in DdCAD-1 phosphorylation. 3.3.10 Summary Cells expressing RasG(G12T) were found to have increased cohesive properties that were mediated by DdCAD-1. These enhanced cohesive properties correlated with a decrease in DdCAD-1 phosphorylation and increased cell surface localization of DdCAD-1. DdCAD-1 phosphorylation decreased during development, at time when DdCAD-1 mediated adhesion increased, consistent with an inverse relationship between DdCAD-1 phosphorylation and DdCAD-1 mediated cell adhesion. rasG null cells were defective in the formation of developmental streams and expressed greatly reduced levels of DdCAD-1. 71 (A) 5.5 — pi 5.5 — p i — ! (B) 1 2 3 4 29-Figure 26. Levels of total and phosphorylated DdCAD-1 in a rasG null and Ax2 control cells. (A) Total protein from a rasG' and Ax2 was fractionated by 2D electrophoresis. Blotted membranes were probed with anti-phosphothreonine antibody to detect phosphoproteins. The section of the 2D immunoblot blot known to contain DdCAD-1 is shown (a = DdCAD-la , b = DdCAD-lb , c - DdCAD-lc a). (B) Total protein from induced Ax2::MB (lane 1), induced Ax2::MB-rasG(G12T) (lane 2), rasG~ (lane 3) and the cadA' (lane 4) cells were fractionated by ID electrophoresis and blotted with a DdCAD-1 specific antibody. Molecular weight (kDa) markers and pi ranges are indicated. 7 2 3.4 Effect of RasG(G12T) on Dictyostelium PKB phosphorylation 3.4.1 Introduction The finding of a decrease in the level of phosphorylated P K B in response to the expression of RasG(G12T) (Figure 27; Table 1) was of considerable interest, since it has been reported that RasG binds to PI3K (Lee et al, 1999) and PI3K is required for the receptor-mediated activation of P K B (Meili et al., 1999), indicating a potential link between RasG and PKB. The activation and phosphorylation of P K B has previously only been studied in developing cells (Meili et al., 1999; Lim et al., 2001). In this study, P K B was identified as a vegetative phosphoprotein that was affected by the expression of RasG(G12T) (Figure 27; Table 1). A function of P K B in aggregating cells is the generation of cell polarity and chemotaxis to cAMP (Meili et al, 1999), and P K B phosphorylation during aggregation occurs in response to stimulation by the chemoattractant cAMP (Lim et al, 2001). This raised the question as to whether P K B was involved in the generation of cell polarity and chemotaxis in vegetative cells and as a test of this hypothesis the stimulation of P K B phosphorylation in response to vegetative chemoattractant folate (Blusch and Nellen, 1994) was investigated. 3.4.2 Enhancement of folate-induced phosphorylation of Dictyostelium PKB in cells expressing RasG(G12T) To investigate the relationship between activated RasG and P K B phosphorylation during vegetative growth, the effect of folate stimulation on the phosphorylation of P K B in Ax2::MB and Ax2::MB-rasG(G12T) cells was determined. Cells were grown in media free of tetracycline for three hrs before being harvested and washed free of HL5 media. Cells were then resuspended in K K 2 buffer and stimulated by the addition of 50 pM folate. Since the activation and phosphorylation of P K B in response to cAMP shows complex kinetics (Meili et al., 1999; Lim et al., 2001), it was possible that P K B phosphorylation in response to folate might be equally complex. In addition, the phosphorlyation of P K B has been previously monitored by ID gel electrophoresis of total proteins samples (Lim et al., 2001) and a myc-tagged version of P K B has been immunoprecipated and shown to react against the phospho-threonine antibody used in this thesis (Lim, 2002). Since ID gel electrophoresis allows the comparison of multiple samples, it was therefore used to analyze the affect of folate stimulation on the levels of phosphorlyated P K B . 73 Ax2::MB Ax2::MB-RasG(G12T) I 54 I • MW I 52 6 . 3 - p l - 6 . 6 6 . 3 - p l - 6 . 6 Figure 27. Representative phosphorylation change in vegetative P K B between A x 2 : : M B and Ax2::MB-rasG"(G12T) cells. The phosphoprotein spot identified as PKB (Table 1) is shown in the center of the image expanded 2D immunoblots. Molecular weight (kDA) and pi ranges are indicated. 74 seconds minutes 0 15 30 45 60 2 4 6 8 10 Ax2::MB Ax2::MB-RasG(G12T) v M ijifMrif tttti n g i M M i n i MNI •49 - 4 9 Figure 28. Folate induced stimulation of P K B phosphorylation. Ax2::MB and Ax2: :MB-rasG(G12T) cells were grown in the absence of tetracycline in HL5 media for three hrs and then resuspended in K K 2 at ~ 5 x 107 cells/ml and stimulated with 50 pM folate. At the indicated times (in minutes), 50 pi aliquots of cells were lysed and protein samples fractionated by ID electrophoresis. Blots were probed with the anti-phosphothreonine antibody as described under Materials and Methods. A representative gel from three independent experiments is shown. Phosphorylated P K B (arrow) and molecular weight markers (kDa) are indicated. 75 An increase in the level of phosphorylated P K B was detected within 15 seconds of the addition of folate in Ax2::MB cells stimulated with 50 uM folate (Figure 28). The level of phosphorylated P K B then decreased. A second increase in the fraction of phosphorylated P K B was also detected. 2 minutes after the addition of folate, and by 6 minutes the level of phosphorylated P K B had reached that observed in the initial peak at 15 seconds. Thus, folate induced a bimodal P K B phosphorylation pattern (Figure 28). Cells expressing activated RasG exhibited increased levels of P K B phosphorylation relative to Ax2::MB cells upon exposure to folate (Figure 28). The bimodal pattern of P K B phosphorylation observed in Ax2::MB cells was not as evident in Ax2::MB-rasG(G12T) cells (Figure 28). Instead, the levels P K B phosphorylation remained high in Ax2::MB-rasG(G12T) cells through the time course of the experiment, suggesting that the presence of RasG(G12T) disrupted the bimodal pattern of activation. These results reveal for the first time that the chemoattractant folate stimulates the phosphorylation of vegetative P K B and that this phosphorylation is affected by levels of RasG(G12T). Prior to folate stimulation the levels of phosphorylated P K B in the Ax2::MB-ra,sG(G12T) cells was higher than the Ax2::MB cells (compare zero time points in Figure 28), suggesting a higher basal level of phosphorylated P K B in the presence of RasG(G12T). This result is opposite to that obtained in the 2D analysis, where phosphorylated P K B is decreased in the Ax2: :MB-rasG(G\2T) cells prior to folate stimulation (Figure 27). This discrepancy in the steady state level of phosphorylated P K B in the activated RasG strain relative to the control strain is potentially due to removal of membrane protein in the 2D analysis, since membranes were pelleted after cell disruption (see Materials and Methods). The ID analysis in Figure 25 therefore might detect phosphorylated P K B present in both the cytosol and the membrane. 3.4.3 Cells expressing RasG(G12T) have more P K B in their membrane fractions Rapid translocation to the Dictyostelium plasma membrane upon receptor stimulation with cAMP has been observed for PH-domain containing proteins, including P K B , C R A C and PhdA (Parent et al., 1998; Meili et al., 1999; Moniakis et al., 2001; Funamoto et al., 2001). Since RasG(G12T) expression stimulates P K B phosphorylation in response to folate, it was possible that it also stimulated its translocation to the plasma membrane. To test this hypothesis, Ax2: :MB and Ax2::rasG(G12T) cells, grown for 3 hrs in the absence of tetracycline, were lysed 76 by filtration through a 5.0 um pore membrane and the lysate centrifuged to separate cell membranes from cytosol (Parent et al., 1998; Lim, 2002). The pellet fraction, along with an un-fractionated lysate sample, was then subjected to immunoblot analysis using a PKB-specific antibody to determine the P K B protein level in the membrane (Figure 29). Although the total P K B levels were virtually identical between the two strains (Figure 29), the pelleted membrane fraction from Ax2::rasG(G12T) cells contained more P K B than Ax2: :MB cells (Figure 29). As a control, the immunoblots were also probed for Actin using an actin-specific antibody and the actin levels were found to be the same for the membrane fractions of the two strains (Figure 29). Although the increase in P K B in the membrane fraction of Ax2::rasG(G12T) cells was only slight, three independent repeats of this experiment gave similar results, with the relative amount of P K B found in the membrane fraction of Ax2: :MB cells being 24 ± 4.5, while in Ax2::rasG(G12T) cells it was found to be 37 ± 3.5 (mean ±sd; n = 3) (Figure 29B). A student's /-test of these differences revealed these changes to be statistically significant (t = 3.95, p = 0.05). If the phosphorylated P K B was primarily membrane bound in RasG(G12T) expressing cells, it would have been lost in Figure 27, thus explaining the differences in basal levels of phosphorylated P K B detected in ID and 2D immunoblot analysis (Figures 27 and 28). 3.4.4 Cell polarity in RasG(G12T) expressing cells The increased levels of membrane bound P K B in RasG(G12T) expressing cells might influence the ability of a cell to generate cell polarity in a chemoattractant gradient, since P K B has been shown to localize to the leading membrane edge of a chemotaxing cell (Meili et al., 1999). P K B translocation has been implicated in generating cell polarity towards a chemotactic signal (Firtel and Chung, 2000). Thus it would be predictive that loss of polarity would inhibit chemotaxis. The ability of RasG(G12T) expressing cells to chemotax to a chemoattractant-filled micropipet (Firtel and Chung, 2000) was therefore determined using time lapse microscopy. When a micropipet containing 25 m M folate was immersed in buffer containing Ax2::MB cells, the movement of cells toward the tip was detectable after 20 minutes (Figure 30) and these cells were polarized towards the micropipet (Figure 30). In contrast, the RasG(G12T) expressing cells were less chemotactic to folate (Figure 30) and exhibited little or no cell polarity. The reduced chemotaxis and cell polarity could be consequences of a RasG(G12T)-mediated increase in membrane bound P K B , since this elevated level might interfer with the specific localization of 77 12 3 4 (A) 4 8 -4 8 -<• P K B o Actin (B) 120 100 Relative so Amount 60 of PKB 40 20 0 Total Membrane Bound Figure 29. P K B membrane localization in response to RasG(G12T) expression. Ax2: :MB (lanes 1 and 3) and Ax2::MB-r<zs'G(G12T) (lanes 2 and 4) cells were grown in tetracycline-free media for three hrs. A portion of these cells were lysed by filtration through a 5.0 um pore membrane and centrifuged at 16,000 x g to provide a membrane pellet fraction. The membrane fraction was resuspended in SDS lysis buffer and a non-fractionated total protein sample was prepared by direct resuspension of cells in SDS lysis buffer. (A) Total protein (lanes 1 and 2) and membrane-only (lanes 3 and 4) samples were fractionated by ID electrophoresis and the blots were probed with PKB-specific (closed arrow) or Actin-specific (open arrow) antibodies. A representative gel from three independent experiments is shown. Molecular weight markers (kDa) are indicated. (B) The immunoblots from three independent experiments ± standard deviations were quantified by densitometric analysis as described under Material and Methods. The values normalized relative to the level of total PKB observed in Ax2::MB cells, arbitrarily set to a value of 100%.(Ax2::MB, white; Ax2::MB-rasG(G12T), dotted). 78 Ax2::MB Om >< 20m 40m ~) * 8 W Ax2::MB-RasG(G12T) Om 20m 40m Figure 30. Chemotaxis of A x 2 : : M B and Ax2::MB-rasG(G12T) cells in a spatial folate gradient. Cells washed free of tetracycline and induced in HL5 media for three hrs were seeded at ~ 4 x 105 cells/cm2 in Nunc dishes. After 30 minutes, the media was replaced with 20% HL5 and a micropipet filled with 25 m M folate inserted into the culture. Cell movements were monitored by time-lapse microscopy over 40 minutes. The star (*) indicates an example of cell polarity towards the spatial folate gradient. Images were taken on an Olympus IX-70 inverted microscope. Bar: 100 um. 7 9 P K B at the leading edge of a chemotactically-stimulated cell, preventing adequate directional sensing (Firtel and Chung, 2000). In addition, the bimodal kinetics of P K B phosphorylation in response to folate for the Ax2::MB cells was largely abolished in the Ax2::MB-rasG(G12T) cells expressing RasG(G12T) and this might adversely affect cell polarity. 3.4.5 Summary RasG was found to regulate both the basal and folate-induced level of P K B phosphorylation, and cells expressing RasG(G12T) contained slightly more membrane bound P K B . In addition, the bimodal kinetics of P K B phosphorylation in response to folate was abolished in Ax2::MB-ra.sG(G12T) cells expressing RasG(G12T). RasG(G12T) cells were less polarized and exhibited less efficient chemotaxis in a spatial folate gradient. Since P K B localization is required for efficient chemotaxis (Meili et al., 1999), these results suggested that RasG may influence chemotaxis through its effect on P K B . 80 3.5 Identification of a protein phosphorylation in response to chemoattractant and its modulation by RasG 3.5.1 Introduction During the course of studies on the phosphorylation of ERK2 in response to cAMP in rasC null cells, the phosphorylation of a second protein of 70 kDa (p70), which peaked at 8 minutes after the addition of cAMP, was detected in wild type cells (C. J. Lim, unpublished observations). p70 was also phosphorylated in response to folate, also peaking at 8 minutes in wild type cells (Figure 31). This stimulation was considerably enhanced in the Ax2: :MB-rasG(G12T) cells and reduced in rasG null cells stimulated with folate (Figure 31). It was unlikely that p70 was a complex containing phosphorylated ERK2, since the kinetics of ERK2 phosphorylation were quite different to those for p70. Since RasG clearly modulated the phosphorylation of p70 (Figure 31), I attempted to identity this novel chemoattractant modulated protein. 3.5.2 Use of narrow-range 2D gel electrophoresis for the resolution of p70 To identify the p70 phosphoprotein, 2D electrophoretic fractionation in combination with immunoblot detection was used as described in Section 3.2. Ax2::MB-ra.sG(G12T) cells were stimulated with folate for 4 minutes, which corresponds to the maximum level of p70 phosphorylation in these cells (Figure 31), lysed and proteins separated by 2D gel electrophoresis, using a pi range of 3-10. This gel was blotted to a PVDF membrane and p70 was detected using the phosophospecific M A P K antibody (Figure 32A). Part of the sample was lysed directly in SDS lysis buffer and fractionated during the 2nd electrophoretic fractionation to provide a direct molecular weight comparison for the 2D fractionated sample. Two major phosphoprotein spots were detected, one of which was phosphorylated ERK2 and the other the p70 phosphoprotein running at an approximate pi of 5.4 (Figure 32A). The degree of resolution of the p70 phosphoprotein was determined by matching this immunoblot to its corresponding brilliant blue G-colloidal stained gel (Figure 32B) as described under Materials and Methods. Unfortunately, the p70 component was not well resolved from other proteins (Figure 32B), making the recovery of uncontaminated p70 impossible. 81 (A) mins: 0 2 4 6 8 10 p70>-pErk2t> (B) Ax2::MB-RasG(G12T) _ _ - -47.5 mins: 0 2 4 6 8 10 12 47.5 (C) Ax2 r a s G " 47.5 Figure 31. Detection of a novel 70 kDa protein phosphorylated in response to folate. Induced Ax2::MB-rasG(G12T) (A), Ax2 (B) and rasG null (C) cells were washed free of HL5 media, resuspended in K K 2 at a density of 5 x 106 cells/ml and starved for 30 minutes. Cells were then washed in K K 2 and resuspended at ~ 5 x 107 cells/ml and stimulated with 50 pM folate. At the indicated times (in minutes), 50 pi aliquots of cells were lysed and fractionated by ID electrophoresis. Blots were probed with a phosophospecific M A P K antibody. A novel 70 kDa phosphoprotein termed p70 (closed arrow), phosphorylated ERK2 (open arrow), and molecular weight markers (kDa) are indicated. 82 (B) 10 C -10 111-73-49-33-28-20-***** •' ' • Figure 32. Fractionation of the folate-stimulated phosphoprotein p70 by 2D gel electrophoresis. Ax2::MB-rasG(G12T) cells, grown in the absence of tetracycline in HL5 media for three hrs, were resuspended in K K 2 at a density of 5 x 106 cells/ml and starved for 30 minutes. Cells were then washed and resuspended in K K 2 at ~ 5 x 107 cells/ml and stimulated with 50 pM folate. After 4 minutes, cell aliquots (250 pg) were fractioned by 2D gel electrophoresis as described under Materials and Methods. (A) An immunoblot probed with phosophospecific M A P K antibody. A 50 pM folate-stimulated protein sample (20 pg) lysed directly in SDS sample buffer was included in the 2nd dimension (C). (B) A corresponding brilliant blue G-colloidal stained gel. The position of p70, molecular weight markers (kDa), and pi ranges are indicated. 83 The p70 phosphoprotein was fractionated by narrowing the pi range of the isoelectric focusing from 3-10 to 5-6, a range encompassing the observed pi of 5.4 for the p70 phosphoprotein (Figure 32A). Immunoblot analysis of these narrow-range 2D gels matched the p70 phosphoprotein spot to a single resolved protein spot (Figure 33). This spot was excised from the brilliant blue G-colloidal stained gel, in-gel digested with trypsin (Wilm et al., 1996) and the extracted peptides were used for peptide mass fingerprint analysis. Matching the obtained mass data against the Dictyostelium database (http://dictybase.org/index.html) resulted in the identification of a leucine-rich repeat protein A (LrrA) (Table 2). When selected peptides from this sample were subsequently subjected to MS/MS analysis to generate peptide sequence data, 3 peptides sequences corresponded to portions the LrrA sequence (Table 2), confirming the identification. Table 2: A list of proteins identified as putative p70 candidates Identified Protein Known/ Predicted Function Mass pi % Peptides Matched by (Da) Sequence MS/MS Analysis Genbank Accession Number LrrA Ras Binding Protein 59807 5.4 LTIDNIPSEIGK GTESIIQWLK V S G L G I Q Q D N E K AAK71315 VatA vacuolar H+-ATPase A subunit 68671 5.4 23 L P C N Y P L L T G Q R V G H N Q L V G E I I R AAB50981 3.5.3 LrrA is a predicted Ras-binding protein, but is not p70 The Dictyostelium IrrA gene sequence has been placed in GenBank (accession # AF388909), but as yet, nothing has been published reporting its characterization. Therefore, database homology searches were preformed to provide clues to IrrA gene function. NCBI blast searches (http://www.ncbi.nlm.nih.gov/BLAST/) revealed LrrA to be homologous to the protein SUR-8, a Ras-binding protein that positively regulates M A P K pathways in Caenorhabditis elegans and mammalian cells (Sieburth et al., 1998; L i et al., 2000). The 29% overall identity LrrA shared with C. elegans SUR-8 (Figure 34A) was primarily due to 19 leucine-rich repeats 84 Figure 33. Fractionation of the folate-stimulated phosphoprotein p70 on a narrow-pi range 2D gel. Ax2::MB-rasG(G12T) cells, grown in the absence of tetracycline in HL5 media for three hrs, were resuspended in K K 2 at a density of 5 x 106 cells/ml and starved for 30 mins. Cells were then washed and resuspended in K K 2 at ~ 5 x 107 cells/ml and stimulated with 50 uM folate. After 4 mins, cell aliquots were fractioned by 2D gel electrophoresis using a narrow range pi of 5-6. The resulting gel was brilliant blue G-colloidal stained and the p70 phosphoprotein was localized as described under Materials and Methods. The arrow indicates the position of p70. Molecular weight markers (kDa) and the pi range are shown. 85 (A) 4 O sua-a iKJvS.N^ofg'aJF j[lf3& a p | | c T " I * N ' R C S IrrA suR-a . 3 7 0 3 # < R - 8 J g G H |1D| soo s i o ( B ) lrr8 lrr3 l r r l 4 lrr5 l r r l 5 lrr7 l r r l 6 l r r l 8 lrr4 lrr6 lrr2 l r r l 7 lrr9 l r r l l l r r l 2 l r r l 3 l r r l 9 IrrlO l r r l 2 o GK KKIJLIJSNM ICL|L|GL|L|STWN|V|< I I |I I I I UDG ASM w| E' G A L K NL :G K l EE! jAS EE L G|L|G I. YE K A |F|G N L SMVS V K L E E W N M L E FjG KL Y N L Q VfFNC SKI iSE |W E FjG D L I G L|T I WDM . QLJSNLRQ| PT|I)GA|L|Q LK A T L I N L Q K L D I J F G M RE|VGNLINLQTLDIJRQN WEWSNIM KINlJS JOT? N N RlLlDlLlS S N E L Q I C D I i K S WD I KIlJSIRNty FA KM KL,QQL . KQ|LJFWSNJN|. K L F Y T . H I & L . F . QjLiQA I . F L . SI . I1.TEL . QlLDDL . CLKSL . Rl lNYI . E I A S L . KM T NOpJUR VHP PtLJQ Y V QUh KLIT M RD[N|PQIJKE I . M|RI V . LJT I D N i T l . THL P B p y i T i SUR-8 Qx X(UGX[3X xg|x x g x g x x g . X@X X L Figure 34. Sequence alignment of Dictyostelium L r r A and C. elegans SUR-8. (A) Alignment of C. elegans SUR-8 and Dictyostelium LrrA, regions of significant similarity are boxed. (B) Alignments of the 19 leucine-rich repeat regions (lrr) of LrrA. Consensus sequence of SUR-8 lrr's is shown for reference. 86 (LRRs) found in both proteins and other SUR-8 family members (Figure 34B) (Kobe and Kajava, 2001). To confirm the identity of LrrA as the p70 phosphoprotein, a c-Myc epitope tag was added to the C-terminus of LrrA and this construct was inserted behind an inducible tetracycline promoter. Ax2 cells were transformed with this expression vector and a clone expressing L r r A M Y C (Ax2 : :MB-Lr rA M Y C ) was isolated. This strain was then stimulated with 50 uM folate for 8 minutes, after which cells were lysed in a non-ionic detergent supplemented with phosphatase inhibitors. If LrrA was identical to p70 it would be expected that immunoprecipitated L r r A M Y C would react with the phosphospecific M A P K antibody in immunoblots. To test this, a monoclonal c-Myc epitope tag antibody was added to the samples to form immunocomplexes with the L r r A M Y C , which were then immunoprecipitated by binding to Protein-A agarose beads. The immunoprecipitated complexes were then solubilized in SDS, separated by SDS-PAGE, blotted onto PDVF membranes, and the membranes immunoblotted with both the c-Myc epitope antibody and phosphospecific M A P K antibody (Figure 35). The addition of c-Myc epitope antibody to the above experiments permitted immunoprecipitation of L r r A M Y C in both stimulated (folate) and mock-stimulated (KK2) cells, but the presence of p70 was not detected in stimulated c-Myc immunoprecipitated samples (Figure 35). Total protein samples from stimulated and mock-stimulated cells, which were directly lysed in SDS lysis buffer, revealed p70 to be phosphorylated in stimulated cells before immunoprecipitation (Figure 35). These data indicated that immunoprecipitated L r r A M Y C did not react with the phosphospecific M A P K antibody. It was subsequently learned that Dr. Wen-Tsan Chang from National Cheng Kung University Medical College had created an IrrA null strain. This strain was obtained from Dr. Chang to determine i f folate-stimulated p70 phosphorylation could still be detected in the absence of LrrA. Protein samples taken from IrrA null cells after stimulation with 50 uM folate were used for immunoblot analysis with the phosphospecific M A P K antibody (Figure 36). p70 phosphorylation was still detectable in this strain, confirming that LrrA was not p70. 87 Total Protein KK2 Folate /> .o P L r r A m y c p70 t > probed with Myc Ab probed with p M A P K A b Figure 35. A test for the reaction of L r r A with the phosphospecific M A P K antibody used to detect p70 phosphorylation. A x 2 : : M B - L r r A M Y C cells induced to express L r r A M Y C were resuspended in K.K2 at a density of 5 x 106 cells/ml and starved for 30 mins. Cells were then washed and resuspended in K K 2 at ~ 5 x 10 cells/ml and either mock-stimulated (KK2) or stimulated (50 pM folate) for 8 mins. After which, cell aliquots were lysed in either 6x SDS lysis buffer (Total Protein) or a non-ionic detergent supplemented with phosphatase inhibitors (KK2 and Folate). L r r A M Y C was immunoprecipitated with the addition of a monoclonal c-Myc antibody (Myc IP) and samples used for 1D immunoblot analysis (lanes 4 and 6). In addition, a mock immunoprecipitation was preformed by adding K K 2 to a set of non-ionic detergent lysed samples (Control). 88 seconds minutes 0 30 45 60 90 Ax2 IrrA' O p 7 0 pErk2 Figure 36. Detection of p70 in folate-stimulated IrrA nulls cells. Ax2 and IrrA null cells were resuspended in K K 2 at a density of 5 x 106 cells/ml and starved for 30 minutes. Cells were then washed and resuspended in K K 2 at ~ 5 x 107 cells/ml and stimulated with 50 uM folate. After the indicated times, cell aliquots were lysed directly in 6x SDS Lysis Buffer. Protein samples were then used for ID immunoblot analysis with a phosphospecific M A P K antibody. Phosphorylated p70 (open arrow), phosphorylated ERK2 (closed arrow), and molecular weight markers (kDa) are indicated. 8 9 3.5.4 Isolation of a protein from the IrrA null that co-migrates with LrrA in wild type cells These results suggested that LrrA was a more abundant component that co-migrated with the p70 phosphoprotein, i.e. p70 and LrrA were not resolved by 2D electrophoresis and p70 was not present in high enough quantities to be identified by MS analysis. The IrrA null strain was therefore subjected to narrow-range 2D electrophoresis as described in section 3.5.3. This analysis revealed the presence of a protein localized in the same position as LrrA in the Ax2::MB-rasG(G12T) cells (Figure 37), indicating these proteins co-fractionated on a 2D gel. Consistent with the idea that LrrA obscured the identification of this component during MS analysis, the protein in the IrrA null strained weakly with silver reagent, whereas LrrA was easily detected by brilliant blue G-colloidal staining (compare Figure 37A to Figure 37B). To determine the identity of the p70 protein from the IrrA null cells, the spot was excised and pooled from 5 silver stained gels to allow adequate material for identification. Pooled spots were in-gel digested with trypsin (Wilm et al., 1996) and the peptides extracted from this digest were used for peptide mass fingerprint analysis and MS/MS analysis as before. The protein was identified as vacuolar H(+)-ATPase subunit A (VatA) (Table 2), a component of the vacuolar ATPase complex of Dictyostelium (Fok et al., 1993; Finbow and Harrison, 1997). 3.5.5 Attempts at confirming that p70 is VatA The contractile vacuole network, of which the vacuolar H(+)-ATPase complex is a major component, has been extensively studied in Dictyostelium (Heuser et al., 1993; Fok et al., 1993; Clarke et al, 2002; Liu et al, 2002). Since the vacuolar H(+)-ATPase subunits are also major components, VatA would be expected to be present in larger amounts than LrrA, rather than the opposite (Figure 37). This is probably also explained by the removal of the membrane fraction prior to the analysis, since VatA is a tightly bound membrane protein. The creation of vatA null strain has been repeatedly attempted without success in Dr. Margaret Clarke's Laboratory at the Oklahoma Medical Research Foundation, leading to the conclusion that VatA is essential for Dictyostelium viability (M. Clarke, personal communication). Dr. Clarke provided an antibody that recognizes VatA in immunoblots, and I attempted to use this antibody to immune precipitate VatA to determine i f the precipitated protein was phosphorylated in response to folate stimulation. However, the antibody did not 90 Figure 37. 2D gel analysis of an IrrA null strain for the presence of p70. Ax2 and IrrA null cells were resuspended in KK2 at a density of 5 x 10 6 cells/ml and starved for 30 mins. Cells were then washed and resuspended in K K 2 at ~ 5 x 10 7 cells/ml and stimulated with 50 uM folate. After 8 mins, cell aliquots were fractioned by 2D gel electrophoresis with a narrow range pi of 5-6. (A) The Ax2 lysates were stained with brilliant blue G-colloidal. (B) The IrrA null lysates were silver stained. The location of the LrrA protein and a protein spot localizing to the same location as LrrA (star) in the IrrA null in shown. Molecular weight markers (kDa) and pi ranges are indicated. 91 immunoprecipitate VatA. Since studies with a vatA null strain were not feasible and the VatA antibody did not immunoprecipitate VatA, attempts were made to overexpress tagged versions of VatA to confirm that this protein was in fact p70. Repeated attempts to express c-Myc tagged VatA fusion proteins (on both the C- and N -terminal ends of VatA) were unsuccessful; no transformants expressing the fusion proteins were obtained even when an inducible promoter was utilized. A strain overexpressing wild-type VatA (vatA-oe) was therefore employed to determine i f VatA overexpression would lead to enhanced p70 phosphorylation. This strain grew very slowly and after a few passages would no longer overexpress VatA. Nevertheless, it was possible to obtain sufficient cells that expressed 2-3 times the normal level of VatA so that the effect of increased VatA levels on p70 phosphorylation could be determined (see Figure 38). vatA-oe and Ax2 control cells were stimulated with 50 pM folate and samples taken at various time points after stimulation were used for immunoblot analysis (Figure 38). In Ax2 control cells, p70 phosphorylation was first detectable at 2 minutes after stimulation, whereas in vatA-oe cells this phosphorylation was detectable after 30 seconds (Figure 38). However, although p70 phosphorylation was detectable earlier in the vatA-oe strain, the increased levels of VatA in the cell did not lead to an increase in the levels of phosphorylated p70 as had been hoped. This experiment provides some evidence that p70 is VatA, in that folate-stimulated p70 phosphorylation occurred earlier in the vatA-oe cells, but there was no correlation between the levels of phosphorylated p70 and the amount of VatA. An alternative approach to try to link p70 and VatA relied on the enrichment of the vacuolar H(+)-ATPase complex in the membrane fraction of Dictyostelium cells (Fok et al, 1993; Nolta et al, 1993; Nolta et al, 1994). Ax2 cells stimulated with 50 pM folate for 8 minutes to maximally phosphorylate p70 were pelleted and resuspended in buffered 0.25 M sucrose (Bush et al, 1994). These cells were broken with a Dounce homogenizer and the percentage of broken cells determined by phase-contrast light microscopy (under these conditions, greater than 95% of the cells were broken). Homogenized samples were subjected to differential centrifugation to generate pellet and supernatant fractions, and these fractions were subsequently used for immunoblot analysis with the VatA-specific and phosphospecific M A P K antibodies (Figure 39). VatA and p70 were similarly enriched in the pelleted fractions (PI and P2), a result consistent with the idea that VatA is p70. 92 Ax2 vat-oe seconds minutes seconds minutes 0 15 30 45 60 2 4 6 8 10 0 15 30 45 60 2 4 6 8 10 73- - * p 7 0 73- <3 VatA Figure 38. VatA overexpression changes the phosphorylation of p70. Ax2 and vatA-oe (B) cells were washed free of HL5 media, resuspended in K K 2 at ~ 5 x 107 cells/ml and stimulated with 50 uM folate. At the indicated times, 50 ul aliquots of cells were lysed, fractionated by ID electrophoresis and blots probed with both a phosphospecific M A P K antibody to detect phosphorylated p70 (closed arrow) and a VatA-specific antibody to determined VatA levels (open arrow). Molecular weight markers (kDa) are indicated. 93 S1 P1 S2 P2 73-73 -<dVatA p70 Figure 39. Subcellular fractionation of VatA and p70. Ax2 cells were stimulated with 50 uM folate for 8 mins, pelleted and resuspended in buffered 0.25 M sucrose. These cells were broken with a Dounce homogenizer and homogenized samples were subjected to differential centrifugation to generate pellet and supernatant fractions (SI = supernatant fraction 1000 g for 5 mins, PI = pellet fraction 1000 g for 5 mins, S2 = supernatant fraction 10,000 g for 10 mins, P2 = pellet fraction 10,000 g for 10 mins). These fractions were used for immunoblot analysis with the VatA-specific (open arrow) and phosphospecific M A P K (closed arrow) antibodies. Molecular weight marker (kDa) is indicated. 9 4 3.5.6 Summary A 70 kDa protein (p70) that was phosphorylated in response to chemoattractant stimulation was detected. Since RasG modified p70 phosphorylation it was thus targeted for identification. 2D immunoblot analysis followed by mass spectrometry identified two previously studied proteins, LrrA and VatA, which co-localized with p70. Myc tagged LrrA was not phosphorylated in response to folate, suggesting that p70 was not LrrA and this was confirmed by the continued phosphorylation of p70 in an IrrA null strain. Eliminating or confirming p70 as VatA was less definitive. Overexpression of VatA resulted in changes in p70 phosphorylation and VatA and p70 co-localized upon cellular fractionation, consistent with the idea the p70 is VatA. However definitive proof of this identification is still lacking. Although outside the scope and time frame of this body of work, further experiments involving the creation of a new antibody for the immunoprecipitation of VatA or analysis of the VatA phosphorylation sites via mass spectrometry (Shou et al., 2002) should provide more direct evidence. 95 4 DISCUSSION 4.1 Inducible expression of activated RasG and its effects on discoidin expression In the fifteen years since the Dictyostelium rasG gene was discovered (Robbins et al., 1989), a number of studies have shown that RasG regulates growth, early development and cytoskeletal functions (Khosla et al, 1996; Tuxworth et al., 1997; Zhang et al., 1999; Khosla et al., 2000; Jaffer et al., 2001), but these studies have not yielded a molecular understanding of the signaling pathways involved. Attempts to identify RasG signaling components, using co-immunoprecipitation and affinity chromatography or restriction enzyme mediated integration to produce genetic suppressors of rasG mutant strains, have proved unsuccessful (M. Khosla and G. Weeks, personal communication). The observation that discoidin gene expression changed in response to the synthesis of an activated form of RasG provided an alternative experimental approach, which was used to search for signaling proteins that changed their phosphorylation state during this response (Figures 10 and 11). In total, 10 putative RasG-regulated components were identified by this method. In addition, the modulation of a chemoattractant-induced phosphorylation by RasG(G12T) was demonstrated. This work provides new molecular insights into how RasG functions. The identification of putative RasG signaling targets depended on the inducible expression of RasG(G12T) and two inducible expression systems were analyzed. Prior to the late 1990's only one inducible expression system was available for Dictyostelium expression, the folate-repressible discoidin I? promoter (Blusch et al., 1992) and this system was exploited to express rasG(G\2T) in Ax2 cells (Khosla et al., 1996). Although this system proved to be useful for earlier studies (Liu et al., 1992; Khosla et al., 1996; Zhang et al., 1999; Jaffer et al., 2001), there was an appreciable basal level of expression in the presence of folate, induction in the absence of folate was slow, expression fluctuated during exponential growth and there were inhibitory effects of RasG(G12T) on expression (Secko et al., 2001). The rnrB promoter was tested as an alternative, since exogenous agents induced rnrB expression (Gaudet et al., 1999). However, strains expressing RasG(G12T) under the control of the rnrB promoter became un-responsive to the mutagenic stimulus after several passages of growth in HL5 media (see Section 3.2.2). Most of the studies reported in this thesis made use of 96 a third inducible expression system based on a tetracycline-repressible tet promoter (Blaauw et al, 2000), which did not suffer from the disadvantages displayed by the other two experimental systems. Induction of RasG(G12T) synthesis from the rnrB and tet promoters was accompanied by repression of discoidin mRNA levels (Figure 5 and 10). In addition, RasG, while not essential for discoidin expression, was necessary for high levels of expression (Figure 6). RasG regulation functioned independently of a number of factors (PSF, C M F , folate) and growth conditions (bacteria and axenic media) that are known to regulate discoidin expression (Blusch et al., 1992; Burdine and Clarke, 1995; Gomer et al., 1991; Wetterauer et al., 1995). However, in all these experiments, Discoidin levels were considerably lower in the rasG null cells than in the wild type cells (Figures 6-9), suggesting that loss of RasG signaling affects some regulator of discoidin expression. Although RasG(G12T) overexpression has been previously shown to reduce the final cell density observed during growth (Khosla et al., 1996), to cause changes in cell shape (Zhang et al., 1999) and to prevent aggregation when nutrients are removed (Khosla et al., 1996), there is no evidence that any of these effects are directly related to the effect on discoidin gene expression. Thus, the data presented in this thesis indicated that RasG had two effects on discoidin expression; it was required to maintain high Discoidin levels but, when present in an activated form at high concentrations, discoidin gene expression was repressed. It is possible, however, that RasG(G12T) acted as a dominant negative form of the protein rather than as an activated form, and thereby inhibited signaling pathways that it does not normally regulate. For example, such a dominant negative effect of the RasG(G12T) protein could function through the sequestration of effector proteins, either through increased binding of RasG effectors or alternatively the binding of the effectors of Dictyostelium Ras proteins other than RasG. However this possibility is unlikely, given the fact that the effects of RasG(G12T) on cell shape and the initiation of aggregation (Khosla et al, 1996; Zhang et al, 1999) are quite different to the phenotypes of the rasG null strain (Tuxworth et al, 1997). For example, rasG null cells do not exhibit the flattened characteristics ofpVEII-rasG(G\2T) transformants and rasG null cells are capable of initiating development (Tuxworth et al, 1997). Furthermore, while rasG null cells and pVEII-rasG(G\2T) transformants both exhibit impaired growth, the nature of the growth inhibition is quite different in the two strains (Khosla et al, 1996; Tuxworth et al, 1997). 97 Nevertheless, it will be important to consider possible alternative approaches for investigating the activation of the RasG pathway to resolve this issue in the future, for example, through the overexpression of wild-type RasG and RasG(G12T) in the rasG null strain or the use of activating stimuli upstream of RasG (Kae et al, 2004). Three other Dictyostelium proteins have been shown to modulate discoidin expression: Ga2, P K A and Dd-STATb (Blusch et al, 1995; Primpke et al, 2000; Zhukovskaya et al, 2004). Ga2 null cells are reduced in their induction of discoidin expression at the onset of starvation after growth on bacteria (Blusch et al, 1995). Cells disrupted in the P K A catalytic subunit (PKA-C) express reduced levels of Discoidin during growth in axenic media, but are still able to induce Discoidin expression in response to PSF and C M F (Primpke et al, 2000). Dd-STATb null cells express higher levels of discoidin mRNA than wild type cells (Zhukovskaya et al, 2004). It is not uncommon in eukaryotes to have multiple pathways acting on a gene, so RasG, Ga2, P K A and Dd-STATb could be in independent pathways. But i f the proteins were linked, epistasis experiments should be able to demonstrate such a linkage. 4.2 Combining RasG activation and functional proteomics to uncover novel phosphoproteins Proteomic techniques provide powerful approaches to study biological problems that were intractable a mere decade ago. A prime example is the recent undertaking to understand biological systems as a whole, i.e. "systems biology", which relies heavily on proteomics, computational biology and genomics (Hood, 2003). Overall the goal of using proteomics has been to better understand biological problems by looking at global changes in cellular proteins (Graves and Haystead, 2002). The study of signal transduction has benefited from the application of proteomic techniques to specific problems and this is well illustrated by the study of growth-promoting signals in mammalian cells. For example, the effect of the mitogen fibroblast growth factor 2 (FGF2) on MCF7 human mammary epithelial cells was analyzed by 2D gel electrophoresis and mass spectrometry (Vercoutter-Edouart et al, 2001). This study monitored changes in protein expression upon FGF2 stimulation, finding four proteins that were up-regulated after 12 hrs, including heat shock protein 90 (HSP90), HSP70, proliferating cell nuclear antigen (PCNA), and the transcriptionally controlled tumor protein (TCTP) (Vercoutter-Edouart et al, 2001). In 98 another study, the MEK1 inhibitor PD98059 was used to inhibit the ERK1/2 pathway in human breast epithelial cells to 10-25% (Seddighzadeh et al., 2000). This 2D electrophoretic analysis detected alterations in the expression of seven proteins upon PD98059 treatment, such as annexin V, HSP80 and cytokeratin 8 (Seddighzadeh et al., 2000). In a similar approach, 12-phorbol 13-myristate acetate (PMA) activation of M A P K in human erytholeukemia K562 cells was combined with selective activation and inhibition of the upstream M A P K components MKK1/2 (Lewis et al, 2000). This analysis detected 25 putative targets of the M K K 1 / 2 - M A P K pathway in PMA-treated cells, of which only five had been previously characterized. The above approaches are useful for studying global protein expression changes in response to various signals, but signal transduction cascades also involve more rapid changes in protein phosphorylation (Hunter, 2000). Therefore, over the last six years, proteomics techniques have been developed to monitor changes in protein phosphorylation in an effort to examine the "phosphoproteome" (Graves and Haystead, 2002). One approach involves the use of 2D immunoblotting with phospho-specific antibodies and mass spectrometry (e.g. Godovac-Zimmermann et al., 1999; Soskic et al., 1999; Yanagida et al., 2000; Yuan and Desiderio, 2003). One notable early example was the identification of proteins that responded to platelet-derived growth factor (PDGF) stimulation of mouse fibroblasts (Soskic et al., 1999). A number of the identified proteins had previously been reported to be involved in the PDGF signaling pathway (for example, ERK1, Akt and Src), but several novel components were also reported (for example, FGR and PTP-2) (Soskic et al., 1999). Immunoprecipitation with phosphotyrosine antibodies, which allows the enrichment of tyrosine phosphorylated proteins before electrophoretic fractionation has also been used (Graves and Haystead, 2002). This approach led to the identification of the tyrosine phosphorylation of Vav-2 in response to EGF stimulation (Pandey et al., 2000) and to the identification of phosphotyrosine proteins involved in colony-stimulating factor-1 stimulation of macrophages (Yeung et al., 1998). Immunoprecipitation has been less successfully applied to the study of phosphothreonine- and phosphoserine-containing proteins due to a lack of phosphoserine and phosphothreonine antibodies capable of immunoprecipitating a wide selection of phosphoproteins (Graves and Haystead, 2002). Other techniques that have been described for the proteomic investigation of protein phosphorylation include: the conversion of phosphoserine residues to biotinylated residues followed by avidin affinity chromatography (Oda et al., 2001), multi-step conversion of phosphoamino acids in 99 protein digests into free sulfhydryl groups and the capture of the derivatized peptides (Zhou et al., 2001), and the use of immobilized metal affinity chromatography (IMAC) based on the affinity of negatively charged phosphate groups for positively charged metal ions (e.g. Fe 3 + or Ga ) (Ficarro et al, 2002). However, these last three techniques have yet to be extensively applied to the study of signaling pathways. In the studies described in this thesis, a functional proteomic approach involving 2D immunoblotting with phosphospecific antibodies and mass spectrometry (Soskic et al., 1999) was used to identify proteins that are potentially involved in RasG signaling. With this approach, it was possible to identify thirteen well-separated vegetative proteins that underwent changes in phosphorylation upon the expression of RasG(G12T), considerably more proteins than are generally identified by more traditional methods, such as immunoprecipitation or yeast-two hybrid analysis. These putative changes in phosphorylation appeared to respond to RasG(G12T) expression, since Ax2::MB and Ax2::MB-rasG(G12T) cells were grown in the presence of tetracycline, conditions in which RasG(G12T) is not noticeably expressed, extracts from these two cells showed similar patterns of protein phosphorylation (D. Secko, unpublished observations). Two of the identified proteins, namely actin and P K B , had been shown previously to be phosphorylated (Jungbluth et al, 1995; Meili et al., 1999), but this is the first report of a change in their phosphorylation state in response to the expression of RasG(G12T). The identification of actin and P K B underlines the potential usefulness of this approach for the detection of downstream signaling components. Since RasG(G12T) expression is known to have multiple effects on the cell that involve more than one downstream signaling pathway (Zhang et al., 1999), it's not surprising that these RasG-responsive phosphoproteins were quite diverse in their predicted function (Table 1 and 2). In addition, these phosphoproteins are not necessarily RasG signaling components and some of the phosphorylations may be end products of a signaling pathway. Of these phosphoproteins, RasGEF-R and DdCAD-1 were found to be absent in their respective null strains (Figure 17 and 18). Similar validation will ultimately be necessary for each of the other proteins, since the possibility cannot be ruled out that each of the identified phosphorylated proteins co-migrates with a more abundant protein species that was not phosphorylated. This potential problem is well exemplified by the attempt at the identification of p70 in Section 3.5. In addition, it will be important to begin to validate the threonine and tyrosine phosphorylation of these proteins, for example, through the analysis of phosphorylation sites via 100 mass spectrometry (Shou et al, 2002), phosphoamino acid analysis, or the immunoprecipation of tagged versions of these proteins. 4.3 The role of Dictyostelium kinases and phosphatases The ultimate goal for identifying phosphoproteins responsive to Ras(G12T) (Table 1) would be to use them to find kinases and phosphatases responsible for the changes in phosphorylation. A number of kinases and phosphatases have been identified in Dictyostelium and it is therefore pertinent to discuss the function of a few of the better-characterized examples that could be regulated by RasG. - Two extracellular signal-regulated kinases (ERKs), also called mitogen-activated protein kinases (MAPKs), have been identified in Dictyostelium. ERK1 was the first to be identified and expression of antisense ERK1 mRNA using the discoidin promoter originally suggested that it was essential for vegetative growth (Gaskins et al., 1994). Subsequently, erkl null cells were created. These cells were viable but showed defects in chemotaxis and development (Sobko et al., 2002). ERK1 reacts with phosphotyrosine antibodies (Gaskins et al, 1994), suggesting it may be regulated in a manner similar to mammalian E R K proteins, which require phosphorylation on threonine/tyrosine residues for activity, although this has yet to be confirmed (Pearson et al., 2001). The second E R K to be identified, ERK2, is required for aggregation, since erk2 null cells do not activate adenylyl cyclase upon receiving a pulse of cAMP and activation of adenylyl cyclase is necessary for aggregation (Segall et al., 1995). erk2 null cells also chemotax poorly to both cAMP and folate, possibly due to defects in polarization (Wang et al., 1998). Based on these phenotypes it is not surprising that ERK2 is phosphorylated and activated in response to both cAMP (Maeda et al., 1996; Kosaka and Pears, 1997) and folate (Maeda et al., 1997; Kosaka and Pears, 1997). ERK2 kinase activity peaks approximately one minute after stimulation and has been correlated with ERK2 phosphorylation (Kosaka et al., 1998). In mammalian cells, E R K proteins are components of M A P K cascades, which include upstream M A P / E R K kinases (MEKs) and M E K kinases (MEKKs) (Pearson et al, 2001). Two such proteins have been identified in Dictyostelium: DdMEK-1 and M E K K a . Disruption of DdMEK-1 results in cells that produce small aggregates and chemotax inefficiently, but in these cells ERK2 is still activated to wild-type levels, indicating that D d M E K l is in a different pathway than ERK2 (Ma et al, 1997). Although, it has been suggested that DdMEK-1 may lie in 101 a pathway leading to ERK1, since the developmental phenotypes of erkl and mek-1 null cells are similar and the overexpression of constitutively active DdMEK-1 in erkl null cells does not alter the erkl phenotype (Sobko et al, 2002). Alternatively, M E K K a may be in a pathway separate from ERK1 and ERK2 that is involved in regulating cell-type patterning and morphogenesis (Chung et al, 1998). Since ERK2, MEK1 and M E K K a cells all have different developmental phenotypes, it would appear that there are at least three M A P K cascades functioning in Dictyostelium, at least during development, but the other components of these cascades have yet to be determined. s. The most extensively studied Dictyostelium kinase is P K A , which is essential for aggregation, but which also functions during other stages of the life cycle (Loomis, 1998; Primpke et al, 2000). It is composed of a single catalytic subunit (PKA-C) associated with a single regulatory subunit (PKA-R), as opposed to two of each subunit as seen in many other organisms (Burki et al, 1991; Mann et al, 1992; Simon et al, 1992). Disruption of P K A - C prevents aggregation (Mann et al, 1992), which has been linked to the loss of preaggregative gene expression (Schulkes and Schaap, 1995). P K A activity is determined by the level of intracellular cAMP, which is regulated by the rate of cAMP synthesis by adenylyl cyclases (Saran et al, 2002) and cAMP degradation by RegA (Shaulsky et al, 1998; Thomason et al, 1998). RegA is part of a two-component system that has been linked to the control of terminal differentiation (Thomason et al, 1999). A pathway involving RegA, ERK2, P K A and potentially RasG has been postulated (Lim et al, 2003). This hypothetical pathway is based on the observation that the disruption of RegA in cells lacking ERK2 restores the ability of these cells to aggregate. This finding suggests that ERK2 inhibits the phosphodiesterase activity of RegA (Shaulsky et al, 1998), leading to the accumulation of cAMP and the activation of P K A (Loomis, 1998). In cells expressing RasG(G12T), there is a reduction in the phosphorylation of ERK2, which could result in insufficient cAMP accumulation and thus decreased P K A activity. The loss of P K A activity could explain the inability of cells expressing RasG(G12T) to aggregate (Khosla et al, 1996). In the above model, ERK2 influences P K A activity, but P K A can also influence ERK2 activity. For example, as discussed in more detail below, the addition of cAMP to wild type cells results in the activation of ERK2, and pka null cells show lower more extended levels of ERK2 activation in response to cAMP and P K A overexpression increases ERK2 activation in response to cAMP (Aubry et al, 1997). In addition, expression of the P K A catalytic 102 subunit in erk2 null cells rescues their aggregation-deficient phenotype, indicating that P K A expression can bypass some functions of ERK2 (Aubry et al, 1997). Although these results suggest a cAMP-regulated pathway that involves P K A and ERK2, the direct phosphorylation of either P K A or ERK2 by other components or each other has yet to be demonstrated. The expression of P K A subunits in rasG mutant strains should help evaluate the role of RasG in the above model. P K B is rapidly phosphorylated and activated upon the reception of a cAMP signal at the cell surface (Meili et al, 1999; Lim et al, 2001). Concurrent with its activation, P K B translocates to the plasma membrane and is localized to the leading edge. The localization of P K B at the leading edge suggests that P K B is important for regulating directional sensing during chemotaxis and, in fact, pkbA null cells lack cell polarity and also move slower in a chemotactic gradient (Meili et al, 1999). In addition, aggregation is delayed in pkbA null cells, and these cells form few aggregation centers and can not aggregate at low cell density (Meili et al, 1999). A role for RasG signal transduction in this process is suggested because RasG(G12T) expression affected both the phosphorylation of P K B and levels of membrane bound P K B (discussed in Section 4.8). PAKa, a member of the Ste/PAK protein kinase family, has been shown to be an in vitro substrate of P K B , being phosphorylated on Thr579. This phosphorylation has been suggested to be required for both P A K a activity and subcellular localization (Chung et al, 2001). Members of the Ste/PAK protein kinase family are regulated by the binding of Rac family members to their CRIB domain (Bokoch et al, 2003). The CRIB domain of PAKa can interact with Dictyostelium Racl-GTP in a yeast two-hybrid assay, but it has yet to be shown directly that any Dictyostelium Rac regulates P A K a (Chung and Firtel, 1999). P A K a is believed to function through effects on myosin II, since paka null cells chemotax poorly and are defective in the assembly of myosin II into the cytoskeleton (Chung and Firtel, 1999). A PKB-related protein, PKBR-1, has also been partially characterized in Dictyostelium. This protein is atypical in that it contains no PH domain and is instead constitutively membrane localized (Meili et al, 2000). PKBR-1 is also activated in response to cAMP through a PI3-kinase independent pathway. Several proteins identified in this thesis were dephosphorylated in response to the expression of RasG(G12T), thus raising the possibility that RasG(G12T) expression activated a phosphatase. Three protein-tyrosine phosphatases (PTP1-3) have been described in Dictyostelium, the mutation of which changes the global wild type pattern of tyrosine 103 phosphorylation (Howard et al, 1992; Howard et al, 1994; Gamper et al, 1996). Spalten, a serine/threonine phosphatase of the PPC2 family, has been shown to be required for cell-type differentiation (Aubry and Firtel, 1998). In addition, the recent identification a Dictyostelium phosphatase and tensin homologue (PTEN) homolog and the creation of a pten null strain have linked this PtdIns(3,4,5)P3/PtdIns(3,4)P2 degrading enzyme to the regulation of cell polarity and chemotaxis (Iijima and Devreotes, 2002). To date, none of these phosphatases have been linked to RasG signaling, but as mentioned above, a necessary next step is to identify both the phosphatases, and kinases, that RasG activates. 4.4 Actin phosphorylation and RasG regulation of the cytoskeleton As mentioned previously, several lines of evidence have pointed to a role for RasG in regulating the actin cytoskeleton and, in particular, F-actin distribution (Tuxworth et al, 1997; Zhang et al., 1999). In mammalian cells, Ras proteins have been implicated in the regulation of the actin cytoskeleton through the activation of Rac (Burridge and Wennerberg, 2004). This activation leads to the activation of P A K and W A V E and subsequently L I M kinases and the Arp2/3 complex respectively. Rac activation is crucial in mammalian cells for the generation of actin-rich lamellipodial protrusions (Burridge and Wennerberg, 2004). In Dictyostelium, 15 homologs of mammalian Rac proteins have been identified and the functions of several of these proteins have been investigated by gene disruption and overexpression studies (Rivero et al., 2001). For example, RacE has been implicated in organization of the cortical cytoskeleton since racE null cells are unable to grow in suspension due to cytokinesis defects (Larochelle et al, 1996). The function of the Rac lA, B and C isoforms have also been investigated through the expression of constitutively active and dominant negative forms of these proteins, and the results of these studies link these proteins to the regulation of F-actin (Palmieri et al., 2000). Two PAKs, P A K a and MIHCK, have been studied in Dictyostelium (Lee et al., 1998; Chung and Firtel, 1999) , and recently an Arp2/3 complex has been cloned (Insall et al., 2001). To date, none of these proteins have been shown to be regulated by Ras proteins. It has been reported previously that RasG(G12T) expression induces tyrosine phosphorylation of a protein with the same apparent molecular weight as actin (Chen and Katz, 2000) . In this thesis, I have identified actin as a RasG-regulated phosphoprotein, confirming the earlier report. In addition, I showed that actin was phosphorylated at more than one site in cells 104 expressing activated RasG by the detection of two spots via 2D electrophoresis (Figure 12 and Table 1). Both of the actin species exhibited increases in tyrosine phosphorylation, suggesting the possibility that two tyrosines are phosphorylated upon the activation of RasG, but the pi shift could also be explained by the phosphorylation of a threonine or serine residue. Induction of actin phosphorylation on Tyr53 in Dictyostelium has been reported to occur upon oxygen depletion, inhibition of ATP stimulation, heat shock, azide treatment and the re-feeding of starved Dictyostelium cells (Schweiger et al., 1992; Howard et al., 1993; Jungbluth et al., 1994; Jungbluth et al., 1995), but a second phosphorylation site has not previously been reported. The phosphorylation of Dictyostelium actin has been correlated with cell-shape changes (Howard et al, 1993; Rebstein et ah, 1993; Chen and Katz, 2000), although the functional significance of this phosphorylation is still not clear. Nevertheless, the phosphorylation of actin is potentially a mode of actin regulation. For example, in Physarum polycephalum, threonine actin phosphorylation in response to dry stress prevents its ability to polymerize (Furuhashi et al., 1998). A kinase, actin-fragmin kinase (AFK), involved in actin phosphorylation in P. polycephalum has been identified (Gettemans et al., 1993). A F K phosphorylates actin when in a complex with Fragmin P and thereby blocks the F-actin capping activity of the actin-Fragmin P complex (Gettemans et al, 1995). In mammalian cells, EGF and insulin have been shown to induce serine phosphorylation of actin (Carrascosa and Wieland, 1985; van Delft et al., 1995). More recently, actin was found to be phosphorylated in opossum kidney cells in response to opioids (Papakonstanti and Stournaras, 2002). This phosphorylation was regulated by PI3K and PAK1 and correlated with the redistribution of filamentous actin (Papakonstanti and Stournaras, 2002). More components involved in Dictyostelium actin phosphorylation need to be discovered before it will become clear how this phosphorylation affects cytoskeletal regulation, and whether RasG is directly involved. 4.5 Which RasGEF is the right RasGEF for RasG? RasGEFs are well established as essential components of Ras signaling pathways (Shields et al., 2000; Wilkins and Insall, 2001) and more than 20 putative RasGEFs have been identified from the Dictyostelium genome sequencing project (Wilkins and Insall, 2001). However, none of these have as yet been shown to act on any of the Ras proteins in vitro. The 105 identification of RasGEF-R as a putative component of the RasG signaling pathway is therefore of considerable interest. A variety of RasGEFs, including yeast Cdc25, human Sosl and human RasGRFl, have been found to be phosphorylated (Gross et al., 1992; Douville and Downward, 1997; Yang et al., 2003) and it has been speculated that phosphorylation can result in both negative (human Sosl) and positive (human RasGRFl) regulation. In fact, in the case of human Sosl, phosphorylation has been suggested as a mechanism for negative feedback of Ras stimulation, given that Ras-regulated components have been implicated in the phosphorylation of Sosl (Douville and Downward, 1997). This negative feedback could potentially ensure tight regulation of Ras activation in response to stimulation. Since RasGEF-R was dephosphorylated in response to activated RasG (Figure 12; Table 1), RasGEF-R could be responsible for the activation of RasG and its de-phosphorylation could be a mechanism for the negative feedback of RasG stimulation. Although the phosphorylation of human Sosl has been suggested to have a negative influence on its RasGEF activity (Douville and Downward, 1997), it has recently been shown to elicit RacGEF activity, leading to actin cytoskeleton remodeling (Sini et al., 2004). If this model holds true for RasG and RasGEF-R, RasG-regulated dephosphorylation of RasGEF-R might result in effects on other Dictyostelium GTPases. Further experimentation is required to determine i f RasGEF-R can act as a specific exchange factor for RasG. However, even i f this were shown to be the case, the lack of an observable mutant phenotype for gefR null cells (R. Insall, unpublished observations) as compared to the complex phenotype for rasG null cells (Tuxworth et al., 1997) suggests that RasGEF-R is functionally redundant. The only other RasGEFs that have been partially characterized are RasGEF-A (also called AleA or Aimless) (Insall et al., 1996) and RasGEF-B (Wilkins et al., 2000a). In the case of aleA null strain, its failure to aggregate is dissimilar from the delay seen in the development of rasG null cells (Insall et al., 1996; Figure 25). Both aleA and rasG nulls cells show impaired chemotaxis, reduced activation of adenylyl cyclase and reduced ERK2 phosphorylation in response to cAMP stimulation (Insall et al., 1996; Tuxworth et al., 1997; Lim, 2002). gefB null cells move rapidly, are impaired in macropinocytosis and exhibit significantly delayed aggregation (Wilkins et al., 2000a). Although rasG null cells exhibit delayed aggregation, they do not exhibit the other properties, so it unlikely that RasGEF-B is the exchange factor for RasG. This complexity highlights the importance of the potential 106 RasG/RasGEF-R relationship, since such relationships have been difficult to elucidate by genetic methods alone. Recently, an assay to measure the activation state of Dictyostelium RasG has been developed (Kae et al., 2004). With this development, it will be important to determine the activation state of RasG in the various gef null strains, although the potential redundancy of Dictoystelium RasGEF function could complicate such an investigation. Alternatively, the detailed analysis of protein phosphorylation in strains lacking genes for RasGEFs, using the techniques described in this thesis, could potentially provide information on the phosphorylation events regulated the actions of these proteins. 4.6 A role for RasG in regulating early developmental adhesion through the dephosphorylation of DdCAD-1 A role for RasG signal transduction affecting early development was suggested by the finding that the expression of RasG(G12T) blocked development (Khosla et al, 1996). The marked effect of activated RasG on discoidin gene expression (Figure 3 and 8; Secko et al., 2001), a known molecular marker of the transition from growth to development (Zeng et al., 2000; Primpke et al., 2000), is consistent with such a role. The results described in Sections 3.3 suggest a new facet to the regulatory role of RasG in regulating early developmental adhesion through the phosphorylation of DdC AD-1. The importance of adhesion during Dictyostelium development was recognized over 35 years ago (Gerisch, 1968) and the adhesion protein DdCAD-1 (also known as contact sites B [csB] and gp24) is one of the first proteins to be expressed at the cell surface at the onset of starvation (Beug et al, 1973; Yang et al., 1997). DdCAD-1 has been implicated in mediating initial side-to-side contacts during early aggregation (Coates and Ffarwood, 2001). A second adhesion molecule, termed gp80 or contact sites A (csA), is induced by cAMP as development proceeds (Faix et al., 1992) and mediates a second type of contact in the aggregation stream (Choi and Siu, 1987). gp80 allows tight adhesion between cells after the initial adhesive events mediated by DdCAD-1. In addition to differences in temporal gene expression, gp80 and DdC AD-1 mediated contacts can also be distinguished by their sensitivity to EDTA, since DdC AD-1 forms EDTA-sensitive contacts and gp80 forms EDTA-insensitive contacts (Coates and Harwood, 2001). 107 Although DdCAD-1 has been long implicated as a major adhesion molecule during early development, the finding that it is phosphorylated in Dictyostelium is novel. Since DdCAD-1 was resolved into at least three spots by 2D electrophoresis, it is likely that it is phosphorylated at more than one site (Figure 12). Although these separated components were detected with a phosphothreonine antibody, the pi shift might be due to phosphorylation on alternative amino acid residues or other modifications. The most abundant DdCAD-1 species, based on intensity by immuno- and silver-staining (s57 or DdCAD-la) , showed a slight, but reproducible increase in phosphorylation upon the expression of RasG(G12T). The second and third DdCAD-1 spots (s56 or DdCAD-lb and s55 or DdCAD-lc) were resolved at lower pi values. DdCAD-lb and DdCAD- lc spots were lower in total protein abundance (as determined by their weaker silver staining intensity) as compared to DdCAD-la . The levels of both the phosphorylated DdCAD-lb and DdCAD- lc species decreased in response to the expression of RasG(G12T) and this data was consistent with an overall decrease in DdCAD-1 phosphorylation in response to activated RasG. This conclusion was supported by ID immunoblot analysis (Figure 23). Since, total DdCAD-1 protein levels were not significantly different between RasG(G12T) expressing and control cells (Figure 16), the changes in DdCAD-1 phosphorylation were not a result of changes inDdCAD-1 levels. DdCAD-1 mediated adhesion is known to be homophilic (e.g. binding is through DdCAD-l/DdCAD-1 interactions) and C a 2 + requiring (Brar and Siu, 1993; Wong et al, 1996). The work reported here suggests that DdCAD-1 may also be regulated by phosphorylation. Initial insight into the effect of phosphorylation on DdCAD-1 function was garnered from the observation that cells expressing RasG(G12T) became cohesive during vegetative growth (Figure 19). Constitutive gp80 overexpression in vegetative cells has been found to induce EDTA-stable contacts in suspension (Faix et al, 1992). RasG(G12T)-induced cohesion was instead EDTA-sensitive and could be blocked with anti-DdCAD-1 antibodies (Figure 20), suggesting that it was DdCAD-1 mediated. The fact that the RasG(G12T) expressing cells appeared more cohesive and contained less phosphorylated DdCAD-1, suggests the possibility that dephosphorylation of DdCAD-1 is involved in mediating DdCAD-1 adherence function. It is important to point out, that in the experiments described in Section 3.3.3 and shown in Figure 20, cell-to-substratum adhesion may also have been affected. In fact, the expression of RasG(G12T) has previously been suggested to increase cell-to-substratum adhesion (Chen and Katz, 2000), 108 and therefore, these experiments are likely assessing a combination of cell-to-cell and cell-to-substratum interactions. Although cells grown on bacteria do not express DdCAD-1 until the onset of starvation (Knecht et al., 1987), axenically grown Ax2 cells express high levels of DdCAD-1 without being adhesive (Figure 13; Sesaki and Siu, 1996). This is presumably due to the sequestration of DdCAD-1 in the cytoplasm of vegetative cells (Brar and Siu, 1993). Cells expressing RasG(G12T) contained rings of DdCAD-1 localized on the cell surface (Figure 19), suggesting the possibility that RasG(G12T) levels regulate the movement of DdCAD-1 to the surface. Transport of DdCAD-1 to the cell surface has been shown to occur via a non-classical pathway involving contractile vacuoles (Sesaki et al, 1997) and a considerable fraction of DdCAD-1 was found in the contractile vacuoles of 12 hr developed cells (Sesaki et al, 1997). Contractile vacuoles containing DdCAD-1 are similar to the DdCAD-1 ring structures observed in RasG(G12T) expressing cells (Figure 21). The location of DdCAD-1 on the surface of developing cells is proposed to allow cells chemotaxing towards cAMP to adhere to one another and enter cellular streams (Sesaki and Siu, 1996). If RasG(G12T) expression enhanced the localization of DdCAD-1 to the surface, one would expect cells expressing RasG(G12T) to be more cohesive; and as a result, potentially form larger cellular streams. In standard in vitro cell cohesion assays, developing RasG(G12T) expressing cells were more cohesive (Figure 22), but, the failure of RasG(G12T) expressing cells to initiate development (Khosla et al., 1996) precluded studying the effect of increased cell surface DdCAD-1 on cellular streaming. Since rasG null cells do differentiate after a delay in initiating aggregation, it was possible to examine their aggregation in more detail. They produced disorganized streams that resembled those of cadA null cells (Figure 25). Since DdCAD-1 has been implicated in the initial formation of cellular streams (Sesaki and Siu, 1996), the streaming phenotypes of rasG, and cadA, null cells may be a result of these cells utilizing alternative adhesion systems during aggregation, a hypothesis consistent with the delay in aggregation and thicker, shorter, streams formed in these strains. rasG null cells also express lower levels of DdCAD-1 (Figure 26) consistent with a role for DdCAD-1 in adherence during streaming. It is interesting that rasG null cells express less DdCAD-1, since RasG(G12T) expression was found not to affect DdCAD-1 protein levels (Figure 16). This difference could be 109 due to the short time period used for expressing RasG(G12T) (e.g. three hrs) and longer induction times may result in increased DdCAD-1 expression. It will be of interest to define the signaling pathway leading from RasG activation to DdCAD-1 dephosphorylation. DdCAD-1 shares some homology with the extracellular domain of metazoan cadherins (Wong et al, 1996), which are involved in cell-cell adhesion and the formation of adherens junctions (Nelson and Nusse, 2004). Metazoan cadherins form complexes with catenins to allow the linking of cadherins to the actin cytoskeleton (Jamora and Fuchs, 2002). These complexes are regulated by phosphorylation. Tyrosine phosphorylation of B-catenin by Src or Fer results in the disruption of the cadherins/B-catenin complexes and decreased adhesion (Lilien et al, 2002; Nelson and Nusse, 2004). It is possible that DdCAD-1 phosphorylation performs a similar function. It should be noted that cadherins/B-catenin complexes are phosphorylated on serine/threonine residues, which results in increased stabilization of the complexes (Lickert et al, 2000). Since DdCAD-1 also appears to be phosphorylated on several residues (Figure 18), its regulation by phosphorylation could be equally complex. Finally, an unidentified transmembrane linker has been postulated to bind to DdCAD-1 (Sesaki et al, 1997), however the proteins that interact with DdCAD-1 are currently not known. Determining the pathway leading from RasG to DdCAD-1 will not only help in our understanding of Dictyostelium development, but could produce insights into the regulation of metazoan cell-cell adhesion. 1 4.7 Effect of RasG(G12T) on PKB phosphorylation It has been suggested that Dictyostelium P K B has two threonine phosphorylation sites (T278 and T435) that are required for receptor-mediated activation of its kinase activity (Meili et al., 1999). P K B activation does not occur in pi3kl/pi3k2 null cells (lacking both PI3K1 and PI3K2), suggesting a requirement for PI3K activity for P K B activation (Meili et al., 1999). Since PI3K is a known downstream effector for activated Ras in mammalian cells (Shields et al., 2000) and RasG is known to bind to PI3K1 and PI3K2 in a yeast two-hybrid assay (Funamoto et al., 2002), it is reasonable to hypothesize that RasG regulates Dictyostelium P K B activity through its activation of PI3K1 or PI3K2. The data presented here provides additional evidence that P K B phosphorylation is RasG-regulated. 110 P K B was detected during the search for vegetative proteins whose phosphorylation was altered in response to the expression of activated RasG (Table 1). P K B was detected with a polyclonal phosphothreonine antibody during the 2D immunoblot analysis, consistent with Meili et a/.'s (1999) postulated phosphorylation of P K B on threonine residues. The 2D analysis revealed that P K B phosphorylation was reduced in an activated RasG strain (Figure 12). This was unexpected, since data from mammalian studies show increased P K B phosphorylation in response to an activated Ras protein (Campbell et al., 1998), and it raised the possibility that in Dictyostelium, RasG negatively regulates P K B . An SH2-domain-containing kinase, SHK1, has recently been identified as a negative regulator of P K B activation in Dictyostelium cells, possibly reducing PtdIns(3,4,5)P3 via the modulation of the activity of either PI3K or PTEN (Moniakis et al., 2001), and RasG might act in a similar manner. However, a more plausible possibility for the reduction in P K B phosphorylation in response to RasG is that the removal of membrane proteins during 2D sample preparation resulted in a loss of membrane-bound phosphorylated P K B . Membrane fractions of cells overexpressing RasG(G12T) contained slightly more P K B than control cells (Figure 29) and more phosphorylated P K B was detected in RasG(G12T) expressing cells analyzed by ID immunoblot analysis (Figures 28). To date, RasG activation of PI3K has not been directly demonstrated for any of the Dictyostelium PI3Ks nor have the levels of PtdIns(3,4,5)P3 in rasG mutant strains been analyzed. It is not clear how PI3K stimulation results in P K B phosphorylation. In mammalian cells, PI3K activation is associated with the translocation of some P K B and phosphoinositide-dependent kinase 1 (PDK1) to the membrane (Anderson et al, 1998; Bellacosa et al, 1998). PDK1 can phosphorylate threonine-308 in the kinase domain of this fraction of P K B and the translocation of PDK1 to the membrane is required for this phosphorylation (Filippa et al., 2000; Vanhaesebroeck and Alessi, 2000). Full P K B activity requires this PDK1 phosphorylation and a secondary phosphorylation on serine-473 (Persad et al., 2001). Although a P D K has yet to be identified in Dictyostelium, the observation that cAMP-stimulated PI3K activation induces the translocation of a fraction of PKB (Meili et al., 1999), suggests that P K B may be translocated and then phosphorylated in response to chemoattractant stimulation. Since RasG(G12T) expressing cells contain more P K B in their membrane fractions (Figure 29), and the translocation of the PKB PH-domain fused to GFP to the membrane in response to cAMP is 111 reduced in rasG nulls cells (R. Firtel, personal communication), it is possible that RasG is regulating the translocation of PKB to the membrane. PKB phosphorylation was also observed to increase upon stimulation by folate (Figure 28), a known chemoattractant in Dictyostelium (Blusch and Nellen, 1994). This phosphorlyation was detected by ID immunoblot analysis of total cell lysates, a method that has successfully used to monitor the phosphorlyation of P K B in response to cAMP (Lim et al., 2001; Lim, 2002). Folate signal reception requires the G proteins GB and Ga4 (Hadwiger et al., 1994; Nebl et al., 2002) and results in the stimulation of ERK2 phosphorylation (Kosaka et al., 1998). The increase observed in P K B phosphorylation upon folate stimulation suggests that P K B may be an additional component involved in the transmission of a folate signal and it would be of interest to determine the ability of pkbA' null cells to chemotax towards folate. Nevertheless, confirmation of the phosphorlyation of P K B in response to folate stimulation, for example through the immunoprecipation of tagged versions of P K B , will be required before initiating such studies. The finding that RasG(G12T) expression increased the levels of P K B phosphorylation in cells stimulated by a chemoattractant indicates that RasG may be involved in the regulation of chemotaxis and the generation of cell polarity. RasG(G12T) expressing cells responded poorly in a spatial gradient of folate and exhibited reduced cell polarity (Figure 30). Since P K B has been implicated in proper orientation and chemotaxis to cAMP (Meili et al., 1999), it is possible that the effects of RasG(G12T) expression on these properties are partially mediated through its role in the translocation and phosphorylation of P K B . Dictyostelium has become an important organism for understanding the mechanisms involved in cell polarity and chemotaxis (Devreotes and Janetopoulos, 2003). In the current model, chemotaxis begins with the receptor-stimulated generation of PtdIns(3,4,5)P3 by the reciprocal regulation of PI3K and PTEN at the leading edge of the cell (Devreotes and Janetopoulos, 2003; Williams and Harwood, 2003). Local concentrations of PtdIns(3,4,5)P3 attract PH-containing proteins, and the formation of actin-filled projections. This localization results in cell polarity and subsequent movement toward a chemoattractant (Devreotes and Janetopoulos, 2003; Williams and Harwood, 2003). The results presented in this thesis suggest the possibility that RasG may be involved in the generation of PtdIns(3,4,5)P3, and thus PH domain binding sites, through the activation of PI3K activity. The generation of PtdIns(3,4,5)P3 in Dictyostelium wild type, pi3kl'/pi3k2'and pten cells in response to cAMP stimulation was 112 recently studied by metabolically labeling cells with P followed by T L C (Huang et al., 2003) and similar experiments on rasG mutants strains would allow investigation of the above possibility. 4.8 Additional targets of RasG regulation The significance of the changes in the phosphorylation of the other proteins identified as responding to RasG(G12T) expression (Tables 1 and 3) were not studied further. Nevertheless, it is anticipated that the cellular activity of these proteins will be affected by the changes in phosphorylation and a brief examination of some of their predicted functions is therefore in order. One of the most intriguing proteins to be identified in this study was FtsZA. This protein is most closely related to the bacterial protein FtsZ (Gilson et al., 2003), a bacterial tubulin homolog that is a component of the "Z ring" structure that constricts upon cell division (Romberg and Levin, 2003). In Dictyostelium, FtsZA and a related protein FtsZB have been implicated in mitochondrial division (Gilson et al., 2003). Neither bacterial FstZ nor Dictyostelium FstZA have been previously shown to be phosphorylated, although the bacterial protein FstA, which binds to FstZ, is phosphorylated in Escherichia coli (Romberg and Levin, 2003). If the RasG-regulated phosphorylation of Dictyostelium FtsZA is confirmed, it will suggest a RasG signaling pathway that regulates mitochondrial division. Another potentially important protein exhibiting changes in phosphorylation in response to activated RasG is the phosphatidylinositol transfer protein 1 (PITP1). In mammalian cells, PITPs are involved in exocytosis of secretory granules, PLC-mediated hydrolysis of PtdIns(4,5)P2, and the PI3K pathway (Cockcroft, 2001). In Dictyostelium, two PITPs, PITP1 and PITP2, have been identified and their ability to stimulate phosphoinositide synthesis has been documented (Swigart et al., 2000). In other organisms, the phosphorylation of PITP has been reported (Snoek et al., 1993; Monks et al., 2001), but this is the first report of the potential phosphorylation of Dictyostelium PITP1. PITPs have been shown to enhance PtdIns(3,4,5)P3 production in human neutrophils (Kular et al., 1997). cAMP stimulation of aggregating Dictyostelium cells has been shown to induce the release of protons and this release is blocked by the vacuolar H +-ATPase inhibitor concanamycin A (Flaadt et al., 2000). This process could be regulated by the phosphorylation of 113 vacuolar H -ATPase components such as VatA, and RasG may be involved in this regulation. However, definitive proof of the phosphorylation of VatA was not obtained, due to the difficulties encountered in trying to express VatA proteins and the fact that VatA appears to be essential (see Section 3.5.5). The antibody used to identify VatA was specific for the doubly phosphorylated T X Y motif of human ERK1/2 (Pearson et al., 2001) and it can be assumed that reactive proteins are also phosphorylated on this motif. VatA does contain a T X Y motif at position 547-549 (with the start codon labeled amino acid position 1), which could be the site of putative phosphorylation. A component of the vacuolar H+-ATPase complex is phosphorylated in another organism (Myers and Forgac, 1993), but this is the first report of a component of vacuolar H+-ATPase complex being phosphorylated in Dictyostelium. New functions continue to be discovered for the vacuolar H +-ATPase complex apart from its ability to pump protons (Nishi and Forgac, 2002). For example, in mammalian cells the overexpression the VatE subunit causes altered actin distribution potentially through it involvement in a mSosl-dependent Racl signaling pathway (Miura et al., 2001) and the 16-kDa proteolipid c subunit of the Vo domain has been suggested to play a role in cell growth through Bl-intergrin binding (Skinner and Wildeman, 1999). If VatA proves to be phosphorylated, it will of interest to define the pathway leading from chemoattractant stimulation to VatA phosphorylation through RasG. 4.9 Emerging themes of the RasG signaling pathway(s) and future prospects The phenotypes of cells overexpressing RasG(G12T) have been difficult to probe. For example, a striking alteration in these cells is an unusual shape suggestive of altered actin regulation (Zhang et al., 1999; Weeks and Spiegelman, 2003). However, as many cell properties are tied to the cytoskeleton, it is difficult to segregate the effects of RasG on actin from its effects on other cellular components. The work presented was initiated to begin to disentangle this problem, by revealing changes in protein phosphorylation, which may have effects on various regulatory networks. The specific changes in phosphorylation described in this thesis provide a first step towards mapping out how signals from RasG penetrate the regulatory networks of the cell. The variety of different proteins whose phosphorylation has been shown to respond to activated RasG is large but not surprising, since RasG is known to affect several independent 114 cellular processes (Zhang et al, 1999; Secko et al, 2001). It has been suggested the mammalian Ras protein, H-Ras, interacts with at least eight effector proteins to thereby potentially eight distinct signaling pathways, which each contain three to eight (known) signaling components (Campbell et al., 1998). The work presented in this thesis suggests that RasG will be no different, and it is thus dangerous at this point to make generalizations. Nevertheless, a body of evidence indicates that there are at least three distinct endpoints of RasG signaling: the actin cytoskeleton, the transition from growth to development and the reception of chemotactic signals leading to cell motility. A l l three are important to growth and differentiation in Dictyostelium, as well as the growth and differentiation of other organisms, and it will be important to understand the role that RasG plays in their regulation. The further identification of proteins lying between RasG and these physiological functions are needed to define how these pathways are being controlled. The approach described in this thesis can also be used as a first step in examining a peculiarity of Dictyostelium, in that this organism has more Ras subfamily proteins than Drosophila or C. elegans (Chubb and Insall, 2001). Thus, for example vegetative Dictyostelium cells contain a minimum of four Ras subfamily proteins (RasB, RasC, RasG and RasS) and each appears to have a specific function. One fundamental question is whether, and to what extent, their signaling pathways interact. Identification of protein phosphorylation changes in response to the overexpression of other Ras proteins should help to answer this question. 115 BIBLIOGRAPHY Anderson, K . E., Coadwell, J., Stephens, L. R. and Hawkins, P. T. (1998) Translocation of PDK-1 to the plasma membrane is important in allowing PDK-1 to activate protein kinase B. 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