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Analysis of the effects of the Rap1 and Rasg genes on dictyostelium discoideum cell morphology Rebstein, Patrick James 1996

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ANALYSIS OF T H E EFFECTS OF T H E RAP1 A N D RASG GENES O N DICTYOSTELIUM  DISCOIDEUM CELL MORPHOLOGY  PATRICK JAMES REBSTEIN  B.Sc, The University of Toronto, 1987 M.Sc, The University of Toronto, 1989  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF T H E REQUIREMENTS FOR T H E DEGREE OF DOCTOR OF PHILOSOPHY in T H E F A C U L T Y OF G R A D U A T E STUDIES (Department of Microbiology and Immunology) We accept this thesis as conforming Jtp^Jjie^required standard  T H E UNIVERSITY OF BRITISH COLUMBIA January 19% © Patrick James Rebstein, 1996  In presenting  this  degree at the  thesis  in partial fulfilment  University of  freely available for reference copying  of  department  this or  publication of  British Columbia, and study.  by this  his  or  her  the  The University of British Columbia Vancouver, Canada  that the  I further agree  representatives.  may be It  thesis for financial gain shall not  Department  requirements  I agree  thesis for scholarly purposes  permission.  DE-6 (2/88)  of  is  for  an  advanced  Library shall make it  that permission for extensive granted  by the  understood  that  be allowed without  head  of  my  copying  or  my written  ABSTRACT  The rapl gene of Dictyostelium discoideum is a member of the ras-gene superfamily of low molecular weight GTPase proteins. The D. discoideum rapl gene is expressed both during growth and development. To further characterize rapl, the sequence and organization of the genomic D N A encoding the rapl gene and 1 kbp of the 5' flanking region was determined. The 5' flanking region could activate expression of the (3-galactosidase reporter gene upon transformation into D. discoideum. To examine the action of the Rapl protein in D. discoideum, the rapl cDNA was expressed under the control of the inducible discoidin promoter. Overexpression of the Rapl protein correlated with the appearance of morphologically aberrant vegetative amebae: cells were extensively spread and flattened with an increase in F-actin staining around the cell periphery. Expression of the rapl cDNA also prevented cell detachment from the substratum upon treatment with azide. When starved D. discoideum amebae are stimulated with HL5 medium, the cells rapidly respond by rounding up. By contrast, the Rapl transformant cells showed a pronounced delay in rounding. Site directed mutagenesis was used to determine the requirements for specific conserved amino acids for these effects caused by overexpression of Rapl. The substitution G10V, predicted to prevent nucleotide binding, and the substitution S17N, predicted to restrict the protein to the GDP-bound state, both abolished the ability of Rapl to affect cell morphology, suggesting that GTP binding is required for Rapl activity. Moreover, the substitution G12V which is predicted to increase the proportion of protein binding GTP, modestly enhanced the ability of Rapl to inhibit the rounding of starved cells after nutrient stimulation. By contrast, substitutions at amino acid 38 in the presumptive effector domain reduced but did not totally abolish the ability of ii  Rapl to affect cell morphology. The substitution T61Q, which impairs the ability of mammalian Rapl to revert the phenotype of Ras transformed mammalian cells, did not alter the effect of Rapl on D. discoideum cell morphology. Thus, although the effects of Rapl on D. discoideum cell morphology appear to be regulated by GTP, the unexpected effects of substitutions at amino acid positions 38 and 61 suggests that the Rapl induced effects may involve different effector and regulatory molecules from that required to revert the phenotype of Ras transformed cells. A D. discoideum transformant expressing activated RasG-G12T protein (RasG-G12T)  had an altered cell morphology similar to that of Rapl  overexpression: the cells becameflattenedand spread with a concomitant distribution of F-actin around the cell periphery. RasG-G12T cells also failed to round up and detach upon exposure to azide. However, expression of activated RasG-G12T resulted in the formation of multinucleate cells whereas Rapl expressing transformants remained mononucleate.  iii  TABLE OF CONTENTS  Abstract  ii  Table of contents  iv  List of Tables  viii  List of Figures  ix  List of Abbreviations  xi  Acknowlegements  xii  Dedication  xiii  Introduction  1  Prologue  1  The Ras gene superfamily  1  Domains of Ras superfamily proteins  3  Regulators and effectors of the Ras superfamily  7  Biological roles of Ras  8  Rapl proteins  10  Mutational analysis of Rapl proteins  12  Regulation of the Rapl proteins  13  Summary  :  Action of Ras superfamily proteins on cell morphology The effects of the Ras superfamily on the cytoskeleton Dictyostelium discoideum  14 15 17 22  Life cycle  22  Signal transduction events during the lifecycle of D. discoideum  24  Vegetative growth, starvation and aggregation  24  Cell differentiation  25 iv  The ras gene superfamily in D. discoideum  27  Cell morphology of D. discoideum  29  Rationale and research objective  34  Materials and Methods  35  Materials  35  D. discoideum growth and differentiation  37  Growth of D. discoideum Development of D. discoideum  37 ..  Determination of cell viability Induction of the discoidin promoter  37 38 38  Analysis of cell morphology  39  Analysis of cell-cell adhesion  40  Flow cytometric analysis  40  Transformation of D. discoideum  40  Dark held, Nomarski andfluorescencemicroscopy  41  Molecular Biology  41  Plasmid DNA preparation  41  Southern blot analysis  42  Preparation of cDNA probes  43  Sequencing  43  Electrophoresis and immunoblotting  43  p-galactosidase assay  44  Isolation of the rapl genomic region  45  Sequence analysis of the rapl genomic region  46  Vector constructions  47  Polymerase chain reactions  52  v  Results  54  The Dictyostelium discoideum rapl gene: isolation of the genomic sequence and characterization of the promoter region Introduction  54 54  Isolation of rapl genomic DNA  54  Nucleotide sequence of the rapl genomic DNA including the 5' region  55  Analysis of the rapl promoter  59  Altered morphology of vegetative amebae induced by increased expression of the Dictyostelium discoideum rapl gene  61  Introduction  61  Effect of Rapl overexpression on cell morphology  61  Time course analysis of Rapl protein levels and cell morphology after induction of the discoidin promoter  66  Localization of F-actin  70  Effects of Rapl expression on growth  70  The response of cells to azide treatment  74  Determination of the number of nuclei in transformed cells  74  Cell motility analysis  '.  Analysis of morphology after HL5 stimulation  74 75  Localization of F-actin in cells treated with HL5 after starvation  79  Pattern of tyrosine phosphorylation of actin after HL5 stimulation  79  Analysis of the erasure response of Rapl cells  82  vi  Identification of conserved residues of the Dictyostelium discoideum Rapl protein required to alter cell morphology Introduction  .  84 84  The effect of mutated rapl genes on cell morphology F-actin distribution in transformed cells Analysis of morphology after HL5 stimulation  84 .92 92  The response of cells to azide treatment  95  Growth and development  95  Activation of the rasG gene alters cell morphology in D. discoideum  99  Introduction  99  The effect of RasG protein on D. discoideum cell morphology  99  The localization of F-actin in D. discoideum cells expressing RasG  100  The response of cells to azide treatment  106  Determination of the number of nuclei in transformed cells  106  Analysis of morphology after HL5 stimulation  Ill  Expression of Rapl protein in RasG-G12T cells and expression of RasG in Rapl cells  114  General Discussion  116  References  137  vii  LIST OF TABLES Table 1. Oligonucleotides used for site directed mutagenesis of the rapl gene  51  Table 2. Erasure of the capacity for rapid developmental recapitulation  83  Table 3. The effect of site directed mutations in the rapl gene on cell morphology  87  Table 4. The proportion of vegetative cells with a flat spread morphology  .  90  Table 5. The percentage of cells remaining adherent after treatment with azide  97  Table 6. Generation times of transformants expressing mutated Rapl proteins  98 ,  Table 7. The proportion of cells with a flat spread morphology  104  Table 8. The proportion of cells remaining adherent after treatment with azide  108  Table 9. The number of nuclei per cell.....  110  Table 10. The effects of conserved amino acid substitutions in Ras and Rapl proteins  ,.•  viii  122  LIST OF FIGURES Figure 1. Alignment and structure assignment for RaplA and Ras  4  Figure 2. GTP cascades controlling cell morphology and bud formation  20  Figure 3. The life cycle of D. discoideum  23  Figure 4. PCR mutagenesis of the rapl gene  50  Figure 5. Southern blot analysis of D. discoideum rapl genomic DNA  56  Figure 6. Genomic organization of the D. discoideum rapl gene  57  Figure 7. Nucleotide sequence encoding the rapl gene  58  Figure 8. f3-galactosidase expression under the control of the 1 kb 5' untranslated region of the rapl gene  60  Figure 9. The pVEII Rapl expression vector  63  Figure 10. Expression of Rapl protein in cells transformed with the pVEII Rapl vector  64  Figure 11. Morphology of vegetative Rapl cells  65  Figure 12. Forward and side light scatter analysis of vegetative Rapl cells  67  Figure 13. Time course analysis of cell morphology after induction of the discoidin promoter  68  Figure 14. Time course analysis of Rapl protein upon induction of the discoidin promoter  69  Figure 15. Localization of F-actin in Rapl cells.  71  Figure 16. Growth of the Rapl transformant  72  Figure 17. Cell-cell adhesion properties of the Rapl cells  73  Figure 18. The effect of treating cells with azide  76  Figure 19. The motility of vegetative cells  77  Figure 20 Effect of HL5 stimulation on Rapl and Ax2 cells  78  Figure 21. Localization of F-actin in starved and HL5 stimulated cells.  80  ix  Figure 22. Protein tyrosine phosphorylation after HL5 stimulation  81  Figure 23. Site directed mutations of conserved amino acids of Rapl...  86  Figure 24. Expression of Rapl protein containing G10V and S17N mutations  88  Figure 25. Cell morphology of transformants expressing mutated Rapl protein  89  Figure 26. Rapl protein expression under inducing conditions  91  Figure 27. Cell morphology and F-actin distribution  93  Figure 28. The response of starved cells to the reintroduction of HL5 medium  '.  94  Figure 29. The percentage of cells responding to the reintroduction of HL5 medium after 5 and 10 minutes  96  Figure 30. Morphology of vegetative cells  101  Figure. 31. Induction of RasG protein expression  102  Figure 32. Forward and side light scatter analysis of RasG-G12T cells.  103  Figure 33. Localization of F-actin in Ras transformed cells  105  Figure 34. The effect of treating cells with azide  107  Figure 35. Nuclear staining of Ras transformed cells  109  Figure 36. The response of starved cells to FIL5 medium  112  Figure 37. The percentage of starved cells that respond to HL5 stimulation  113  Figure 38. Expression of Rapl protein in RasG-G12T cells and expression of RasG in Rapl cells  115  Figure 39. Comparison of azide treatment with the HL5 stimulation assay  •  x  •  ••— 128  LIST OF ABBREVIATIONS BSA  bovine serum albumin  CTAB  cetyl trimethyl ammonium bromide  ECL  enhanced chemiluminscence  EGF  epidermal growth factor  FITC-phalloidin  fluorescein isothiocyanate-phalloidin  GAP  GTPase activating protein  GTPyS  guanosine 5'-(3-0-thio) triphosphate  NGF  nerve growth factor  PBS  phosphate buffered saline  PEG  polyethylene glycol  PDGF  platelet-derived growth factor  RCF  relative centrifugal force  SDS-PAGE  sodium dodecylsulfate polyacrylamide gel electrophoresis  TBS  Tris buffered saline  Single letter code for amino acids: A, Alanine; R, Arginine; D, Aspartic acid; C, Cysteine; Q, Glutamine; E, Glutamic acid; G, Glycine; H, Histidine; I, Isoleucine; M, Methionine; F, Phenylalanine; P, Proline; S, Serine; T, Threonine; W, Tryptophan; Y, Tyrosine; V, Valine.  xi  ACKNOWLEDGMENTS I thank Wolfgang Nellen for the pVEII vector; Meenal Khosla for assistance in the vector construction and the RasG and RasG-Gl2T strains; Jasmine Cohen for assistance with sequencing and the p-galactosidase assays; and Sharon Louis for the G10V and G12V mutated rapl cDNAs and helpful discussion.  I thank the Gerish laboratory for performing the cell-cell  adhesion analysis and the cell motility experiments.  I thank Michael  Schleicher, Matt Springer and Bruce Patterson for their helpful discussions on the effects of azide on D. discoideum. I thank Dennis Dixon for helpful discussions and assistance with the microscopy. I thank Dr. Frank Tufaro and Mr. Mike Weiss for letting me use their microscopes. I thank my committee members: Drs. Beatty, Jefferies, and Snutch for their assistance. I thank all members of Dr. Week's lab past and present who gave their advice and support. Finally I thank my supervisors Drs. Gerry Weeks and George Spiegelman for their guidance and efforts to make me think critically. I was a recipient of a Steve Fonyo Award from the National Cancer Institute of Canada and a University Graduate Fellowship. The research was supported by grants from the National Cancer Institute of Canada, the Medical Research Council and the BC Health Research Foundation.  xii  DEDICATION  To Karen and Rachael.  INTRODUCTION  Prologue The primary objective of this thesis was to analyze the effects of expressing the ras-related rapl gene on cell morphology in Dictyostelium discoideum. In addition, the effects of expression of the rasG gene on cell morphology were also analyzed. In the Introduction, I shall initially review the literature on the ras gene superfamily focusing primarily on the well understood Ras proteins which have been extensively characterized both structurally and functionally. I will also review the literature on the rapl genes and other relevantras-relatedgenes. I will place a special emphasis on the roles that members of the ras gene superfamily play in modulating and regulating cell morphology. I will then discuss D. discoideum, focusing on the regulation of cell morphology during both growth and development before summarizing what is known about the ras andras-relatedgenes.  The Ras gene superfamily  The ras superfamily can be subdivided into three major groups that consist of the ras, rho and rab sub-families, based on the degree of shared amino acid conservation and in some cases on the protein function (Valencia et al, 1991; Kahn et al, 1992). More than 50 members of the mammalian ras superfamily have been identified to date. The ras sub-family includes the proto-oncogenes Ki-ras, H-ras and N-ras and the genes encoding the closely related proteins R-Ras and TC21 which share 55% identity with Ras (Cox et al., 1994; Graham et al, 1994) and the rap genes which encode proteins that  share approximately 50% amino acid identity with Ras (Bokoch, 1993; Noda, 1993). The roles of the Rapl proteins is not fully understood but Rapl is capable of antagonizing transformation by Ras in some mammalian cells (Kitayama et al, 1989). The functions of the Rapl proteins will be discussed in more detail below. The ras sub-family of genes encodes proteins which are highly conserved throughout evolution. For example, the D. discoideum RasG protein has 69% conserved amino acid identity relative to H-ras (Robbins et al, 1989), while the D. discoideum Rapl protein is 73% identical to its human counterpart RaplA (Robbins et al., 1990), and the D. discoideum Ras and Rapl proteins are more highly related to the human Rapl and Rapl proteins than they are to the D. discoideum Rab and Rho proteins (Daniel et al., 1993a; Bush et al, 1993a). The identification of the presence of the same sub-families of the ras superfamily in divergent species likely represents some degree of conservation of function (reviewed by Valencia et al, 1991). In some cases, the ability to function equivalently has been demonstrated experimentally. For example, it has been shown that the mammalian H-ras gene can compliment ras deficient S. pombe and S. cerevisiae (Nadin-Davis etal, 1986; Kataoka et al, 1985; DeFeo-Jones et al, 1985). The rab gene sub-family members encode proteins which share approximately 30% amino acid homology with the Ras proteins (Rothman and Orci, 1992). Proteins in this sub-family have been shown to be required for vesicle transport between organelles in cells and for the production of synaptic vesicles (Rothman and Orci, 1992; Sudhof, 1995). The rho gene subfamily encodes Rho, Rac and Cdc42 proteins which all share approximately 30% amino acid identity with Ras proteins (Nobes and Hall, 1994). Rho, Rac and Cdc42Hs are capable of inducing the formation of actin stress fibers,  membrane ruffles and filopodia, respectively (Ridley and Hall, 1992; Ridley et al, 1992; Kozma et al, 1995; Nobes and Hall, 1995).  Domains of Ras superfamily proteins  The Ras superfamily of proteins bind and hydrolyze GTP to GDP and it has been proposed that they act as molecular switches regulated by their nucleotide binding state (Bourne et al, 1990). The structures of the Ras and Rapl proteins have been determined by X-ray crystallography The structures, as determined by X-ray crystallography, of the two proteins are very similar (see Fig. 1 for an alignment of the structures of Ras and Rapl), suggesting a conserved mechanism for binding and hydrolysis of GTP (De Vos et al., 1988; Pai et al., 1990; Nassar et al., 1995). The amino acids required for binding and hydrolyzing GTP are located in 4 domains that are well conserved throughout the Ras superfamily and in additionalflankingamino acids conserved within the Ras sub-family (Fig. 1). These domains have been identified by sequence comparison, analysis of mutated proteins and from the crystal structure of Ras and Ras related proteins (Pai et al, 1990; De Vos et al, 1988; Nassar et al, 1995). The a subunits of the heterotrimeric signal transducing G proteins and the bacterial elongation factor Tu also share limited amino acid homology in these regions reflecting a conservation of GTP binding domains across a broad range of proteins (Bourne et al, 1990). In the Ras sub-family these domains consist of residues 10-17; 53-62; 112-119 and 144-146, respectively (Bourne et al, 1991). (See Figure 1 for conservation of these domains between Ras and Rapl). The first domain (10-17) forms bonds with the a-and (3-phosphates of GTP or GDP; the second domain (53-62) forms a hydrogen bond with the yphosphate of GTP; and the third domain (112-119) forms hydrogen bonds  Res  M l E Y K L V V V G A G G V G K S A L T l O l l O N H F V O E  Y D P T I E O S Y  H K O V V I O G E T C L L 0 1  Rop  M R E Y K L V V l G S G G V G K S A L T V O f V O G  Y O P T I E O S Y  R K O V r v o C O O C M L E I  IF V [ K  i. x > i ).>. x >, y i. J i x n 56  L D T A G O E E Y S A M R D O Y M R T G C G F L C V r A I N N T K S F L D I H O Y B E Q I K R V K O S D D V P M V , L D T A G T E O r T A M R O L Y M K N G O G F A L V Y S I T A O S T F N O L O D L R E O I L R V K O T C O V P M I  56  P5  B  J6  c4  ,,3  L V G N K C O L - A A R T V E S R Q A Q D L A R S Y G -  113  L V G N K C 0 L C D E R V V G K C Q G G" N L A R 0 w C N C A f L E S S A K S K  I PY 1 E T S A K  T ' R C G V f O A F Y T L V R c IN V N L  I ROH •  I F Y O L V R O 1NR -  RAP1_DICDI RAPA_HUMAN RASK HUMAN  MPLREFKIWLGSGGVGKSALTVQFVQGIFVEKYDPTIEDSYRKQVEVDSNQCML M - - REYKLVVLGSGGVGKSALTVQFVQGIFVEKYDPTIEDSYRKQVEVDCQQCML MT- -EYKLVWGAGGVGKSALTIQLIQNHFVDEYDPTIEDSYRKQVVIDGETCLL  RAP1_DICDI RAPA_HUMAN RASK HUMAN  EILDTAGTEQFTAMRDLYMKNGQGFVLVYSIISNSTFNELrPDLREQILRVKDCED 110 EILDTAGTEQFTAMRDLYMKNGQGFALVYSITAQSTFNDLQDLREQILRVKDTED 108 DILDTAGQEEYSAMRDQYMRTGEGFLCVFAINNTKS FEDIHHYREQIKRVKDSED 108 ****** *__**** * * _ * * * * * * * * * **** _ *  RAP1_DICDI RAPA_HUMAN RASK HUMAN  VPMVLVGNKCDLHDQRVISTEQGEEIiARKFGDCYFLEASAKWKVNVEQIFYNLIR VPMILVGNKCDLEDERVVGKEQGQNLARQWCNCAFLESS AKSKINVNEIFYDLVR VPMVX.VGNKCDLPS-RTVDTKQAQDLARSYGI -PFIETSAKTRQRVEDAFYTLVR  RAP1_DICDI RAPA_HUMAN RASK HUMAN  QIN--RKNPVGPPSKAK SKCALL QIN—RKTPVEKKKPKK KSCLLL EIRQYRMKKLNSSDDGTQGCMGLP-CVVM  —  *  *_*_**_*_*********_*__*_  ***********  _ * . . . . *.  * * _ * * * * * * * * * * * * *  **  ..*.***..  55 53 53  _ *' _ * . *  *. .  * * .r**" . *  165 163 161  186 184 189  Figure 1. Ahgnment and structure assignment for Rapl A and Ras A) This figure is modified from (Nassar et al, 1995). p sheets (open arrows)\ and a helixes (spiral bars) for H Ras (Ras) and human Rapl A (Rap). Residues found in all GTP-binding proteins are underlined; effector residues 32-40 are bracketed. B) Alignment of D. discoideum Rapl (RAP1 DICDI) with human Rapl A (RAPA HUMAN) and K-Ras (RASK HUMAN)  with both the guanine ring and the first domain. The fourth domain (144146) is somewhat variable and indirectly interacts with the  guanine  nucleotide by stabilizing the third domain. A n additional domain, the core effector domain (extending from amino acids 32-40), is conserved in the Ras sub-family but is not conserved between members of the Ras superfamily with the exception of threonine 35 which is required to coordinate a Mg2+ ion. Mutations which disrupt this domain abolish the transforming ability of Ras (McCormick, 1994; Marshall, 1993). It has been proposed that this domain interacts with downstream effector molecules such as Raf (Avruch et al, 1994). Comparison of the crystal structure of Ras-GDP with that of Ras-GTP has revealed differences i n the effector region amino acids 32-40 and as well as in another external loop region from amino acids 60-76. It has been hypothesized that changes in the structure of these two loops are involved in the transduction of a signal to the effector protein (Brunger et al, 1990; Milburn et al, 1990). Although the effector domain is conserved in all the Ras sub-family proteins, they do not function equivalently (Bourne et al, 1991). Ral cannot transform cells (Feig and Emkey, 1993); TC21 can transform cells in a manner equivalent to Ras; R-Ras can transform cells but does not cause a characteristic transformed cell morphology while Rapl can antagonize transformation by Ras (Graham et al, 1994; Bokoch, 1993; Cox et al, 1994; Noda, 1993). These results suggest that additional effector specificity lies outside this region. Mutagenesis studies have revealed additional requirements within the extended effector domain of Ras, which encompasses amino acids 25-46 (NurE-Kamal et al., 1992). Rapl and R-Ras diverge from Ras within this region, which possibly explains their differing effects in transforming cells. In the Rho and Rab sub-families there are amino acid substitutions within the core  effector domain and there is evidence that these proteins interact w i t h quite different effector proteins (reviewed i n Nobes and H a l l , 1994; Chant and Stowers, 1995) A n additional small domain is present at the carboxyl terminal of all Ras-related proteins. This domain is subject to post translational processing i n Ras, resulting i n removal of the terminal 3 amino acids, followed by addition of a farnesyl isoprenyl group and carboxymethylation of the processed terminal cysteine (Hancock et al., 1989; Clarke, 1992). This domain w i t h its subsequent modification is required for the attachment of Ras to the inner leaflet of the plasma membrane and disruption of this modification blocks the ability of Ras to transform cells (Guierrez et al., 1989; Williamsen et al., 1984). Additional features i n each Ras protein also contribute to specify its intracellular location, for example, a sequence rich in lysine near the carboxyl terminus i n K-Ras and a palmitoylation modification near the C terminus i n H-Ras and N-Ras are also required for localization to the inner leaflet of the plasma membrane (Clarke, 1992). The post translational processing of R a p l proteins is similar to of Ras, but a geranylgeranyl moiety is added rather than a farnesyl group (Buss et ai, 1991). However, the location of R a p l i n the cell, presumably specified i n part by this modification, has not been conclusively established.  In platelets, R a p l has been reported to be localized i n the  cytoplasmic fraction and to be translocated to a cytoskeletal fraction upon thrombin activation of platelets, suggesting that its location i n the cell is subject to additional regulation (Fischer et al., 1990).  In contrast, i n  neutrophils, R a p l A has been reported to bind to cytochrome b558 of N A D P H oxidase and this binding is inhibited by R a p l A phosphorylation (Bokoch et ah, 1991). T w o conflicting reports have localized R a p l to the G o l g i and the late endocytic compartments in fibroblasts (Beranger et al, 1991; Pizon et al,  1994) . In addition, chimeric Ras proteins containing Rapl amino acids from position 66 or position 111 to the carboxyl terminal are still capable of transforming cells (Buss et al., 1991; Zhang et al., 1991), implying that some Rapl protein might also be located at the inner leaflet of the plasma membrane.  Regulators and effectors of the Ras superfamily  The binding and hydrolysis of GTP by Ras is regulated by other protein components, which in some cases also serve to transmit information. The switch from the GDP bound state to the GTP bound state is mediated by guanine nucleotide exchange factors which stimulate the dissociation of GDP from Ras (reviewed in Boguski and McCormick, 1993). The exchange factors function immediately upstream of Ras and are linked to receptors via adapter proteins such as GRB2 and She (reviewed in Schlessinger, 1994; Pawson, 1995) . Several Ras exchange factors have been identified, possibly linking Ras to a different receptors (Boguski and McCormick, 1993). Elucidation of the effector molecule downstream of Ras had been an elusive goal, but recently three effectors which interact directly with Ras have been identified in mammalian cells: Raf, phosphatidylinositol-3-OH kinase (Pl-kinase) and mitogen-activated protein kinase kinase kinase (MEKK1) (Warne et al; 1993; Moodie et al, 1993; Koide et al, 1993; Zhang et al., 1993; Viciana et al, 1994; Russell et al, 1995). Raf and MEKK1 transduce a signal by activating the MAP kinase cascade while Pl-kinase phosphorylates phosphoinositides at the 3' position (reviewed in Blumer and Johnson, 1994; Cantley etal, 1991).  Ras signaling terminates when GTP is hydrolyzed to GDP. The proteins GAP and NFI enhance the rate of GTP hydrolysis (Trahey and McCormick, 1987; Martin et al, 1990). It is possible that GAP, in addition to downregulating Ras, may also function as an effector molecule.  GAP is  required for Ras dependent inhibition of atrial muscarinic K+ channels and GAP is capable of altering cytoskeletal structure and cell adhesion in a manner independent of Ras (Yatani et al., 1990; Martin er: al, 1992; McGlade et al, 1993). However the mechanism whereby GAP causes these effects is not clear. Numerous exchange factors and GAPs which regulate the guanine nucleotide binding and hydrolysis of other Ras superfamily proteins have been identified (Boguski and McCormick, 1993), suggesting that the regulation of these proteins is similar to that for Ras. Biochemical analysis has revealed an additional level of complexity as some regulatory proteins appear to be able to interact with more than one Ras superfamily member. For example, the exchange factor, smgGDS, interacts functionally in vitro with Ras, RhoA and Rapl (Mizuno et al, 1991) while RalGDS interacts with both Ral and Ras in vitro (Hofer etal, 1994). Furthermore, R-Ras interacts with Ras-GAP, NFI and also Raf (Spaargaren et al, 1994; Rey et al, 1994). Although, the significance of these interactions remains to be demonstrated in vivo, the potential clearly exists for complex interacting networks of Ras related proteins.  Biological roles of Ras  Activated ras genes were first identified in viral or tumor cell D N A based on their ability to transform NIH3T3 cells (reviewed by Barbacid, 1987;  Der, 1989; Lowy and Willumsen, 1993). Mutations at codons 12,13 and 61 were identified in the three  ras  genes,  H-ras,  Ki-ras  and N-ras  ,  in many  human tumors (reviewed by Bos, 1989). Mutated ras genes were identified in 80% of pancreatic tumors, 50% of colon carcinomas, 30% of non-small cell lung carcinoma and 20% of melanomas. Different tumor types were usually found to contain mutations in specific ras genes. For example, pancreatic and non-small cell lung tumors had mutations in  Ki-ras,  while melanomas  contained mutations in N-ras. Microinjection of activated Ras proteins in NIH 3T3 cells caused cell proliferation and resulted in a transformed cell appearance (Feramisco et al, 1984; Stacey and Kung, 1984; Mulcahy et ah, 1985). Taken together, these observations strongly implicated mutated ras genes in the process of human neoplasia. This hypothesis was confirmed experimentally by the observation that transgenic mice expressing an activated  N-ras  gene from the MMTV promoter elicited tumors in the  mammary and salivary glands (Mangues et al, 1990; Mangues et al, 1992). In vitro,  activated Ras protein transforms cells and bypasses the  requirements for serum and anchorage dependent growth (Hall et al, 1983; Spandidos and Wilkie, 1984; Paterson et al, 1987) while microinjection of Ras-GTP stimulates DNA synthesis in fibroblast cells (Stacey and Kung, 1984; Feramisco et al, 1984). These studies suggest a role for Ras in cell proliferation. Conversely, microinjection of Ras specific antibody blocks the mitogenic activity of serum and growth factors (Mulcahy et al., 1985). Furthermore, the proportion of Ras-GTP increases after treatment of cells with serum or growth factors such as PDGF or EGF, suggesting that Ras mediates the response to these factors, and is consistent with the idea that binding of GTP results in an active form of the protein (Satoh et al, 1990b; SaXoh etal.,  1990a).  A role for Ras proteins in differentiation and development has also been well documented. PC12 pheochromocytoma cells develop into neural type cells upon treatments with NGF in a process which is dependent on Ras (Bar-Sagi and Feramisco, 1985; Satoh et al, 1987). In addition, transfection of Ras can induce differentiation of 3T3-L1 fibroblasts into adipocytes (Benito et al, 1991). A role for Ras in development has also been identified by genetic analysis in C. elegans and D. melanogaster (Beitel et al., 1990; Simon et al., 1991; Lu etal., 1993). In C. elegans, activation of Ras produces a multivulval phenotype while loss of Ras activity leads to a vulvaless phenotype, again consistent with a switching function for Ras in a signaling pathway. In D. melanogaster, activation of Ras disrupts normal cell fate specification in the compound eye and when microinjected into embryos, disrupts the terminal cell fates of posterior cells (Simon et al, 1991; Lu et al, 1993). Although Ras proteins have been implicated in both proliferation and diverse developmental processes, it appears that in many cases that Ras transduces a signal via a similar pathway that is conserved through evolution.  The Ras effector, Raf, is required for the stimulation of  proliferation by Ras and also for the developmental events mediated by Ras in C. elegans and D. melanogaster  (Moodie and Wolfman, 1994).  Furthermore, the subsequent activation of a MAP kinase cascade by Raf was also found to be an evolutionarily conserved feature.  Rapl proteins  The rapl genes (Pizon et al, 1988a) (also called Krev-1 (Kitayama et al, 1989) and smg p21 (Kawata et al, 1988)) encode low molecular weight GTPase proteins and have been identified in organisms as diverse as S. cerevisiae and  mammals (Schejter and Shilo, 1985; Pizon et al, 1988a; Pizon et al, 1988b; Kawata et al, 1988; Bender and Pringle, 1989; Robbins et al, 1990; Hong et al, 1990; Hariharan et al, 1991). A defining characteristic of Rapl proteins is a conserved threonine at position 61 which is a conserved glutamine in Ras genes (Bourne et al, 1991; Noda, 1993). In humans there are two rapl genes, rapl A and raplB, which encode proteins that are 95% identical to each other (Pizon et al, 1988a; Pizon et al, 1988b). In addition, a related pair of genes encode the Rap2A and Rap2B proteins, which are approximately 60% identical to Rapl A and RaplB (Pizon et al, 1988a; Farrel et al, 1990; Ohmstede et al, 1990).  The Rap2A protein cannot antagonize Ras-induced  transformation (Jimenez et al, 1991). A raplA cDNA, Krev-l, was isolated by its ability to suppress the transformed phenotype of Ki-ras transformed NIH 3T3 cells (Kitayama et al, 1989). Flattened, more adherent cells with reduced tumorigenicity were isolated following transfection with the rapl A cDNA. Rapl proteins antagonize Ras function in a variety of in vitro assays. They inhibit Rasdependent activation of MAP kinases, ERK1 and ERK2 (Cook et al, 1993); NRas induced inhibition of M2-muscarinic receptor coupled K channels in +  heart (Yatani et al, 1991) and H-Ras induced activation of germinal vesicle breakdown in Xenopus laevis oocytes (Yatani et al, 1991; Campa et al, 1991). In view of the conservation of core effector domain between Rapl and Ras proteins (Pizon et al, 1988a; Kitayama et al, 1989) (Fig. 1), it has been proposed that Rapl directly competes with Ras for a downstream effector proteins such as Raf or possibly Ras-GAP, both of which have been shown to interact directly with Rapl (Freeh et al, 1990; Zhang et al, 1993). The Rapl proteins do not appear to act solely as antagonists of Ras since they appear to have different roles in various cell types, and they may signal  through as yet unknown effector molecules (Kitayama et al, 1989; Yoshida et al., 1992). In platelets, RaplB is activated by agents that elevate intracellular cAMP (Altschuler et al., 1995). Rapl has also been implicated in the oxidative burst in B lymphocytes (Maly et  ah,  1994). In S.  cerevisiae,  the rapl gene  homologue RSR1/BUD1 is required for bud site localization, a process proposed to involve the orientation and localization of the actin cytoskeleton in the cell (Bender and Pringle, 1989; Chant and Herskowitz, 1991). Finally, Rapl can elicit some effects that are similar to those of Ras. In Swiss 3T3 cells, microinjection of RaplB in the presence of insulin partially mimics the effect of Ras, causing DNA synthesis and membrane ruffling (Yoshida et al, 1992) and in S.  pombe,  both the human RaplA and RaplB cDNAs were isolated as  genes capable of suppressing the morphological and sporulation defects in a rasl  mutant strain (Xu et al., 1990).  Mutational analysis of Rapl proteins Mutagenesis and domain exchanges have been used to compare the Rapl and Ras proteins (Zhang et al, 1990; Kitayama et al, 1990; Zhang et al, 1991).  Using cell transformation as a criteria for Ras activity and  transformation suppression as a criteria for Rapl activity, domain exchange experiments located the amino acids which specified Ras or Rapl activity to a limited region flanking the core effector domain (Zhang et al, 1990). Further analysis showed that simply substituting Rapl amino acids 26, 27, 30, 31 and 45 with the corresponding residues from Ras was sufficient to generate a protein with Ras-like properties in both mammalian and yeast biological assays (Marshall et al., 1991). These results suggests that effector specificity is encoded in a small number of amino acids in extended effector region flanking the core effector.  Mutational analysis of Rapl amino acids that are conserved between Ras and Rapl suggested a similar relationship between structure and function for the two proteins. Mutations that activate Ras cause enhanced tumor suppression by Rapl while mutations in the core effector region that block transformation by Ras cause attenuated tumor suppression by Rapl (Kitayama et ah, 1990). The requirement for GTP binding for Rapl activity was also evaluated by microinjecting RaplB into Swiss 3T3 cells in the presence of insulin (Yoshida et al, 1992).  When bound to the non  hydrolysable GTP analog GTPyS, RaplB induced DNA synthesis and membrane ruffling, whereas GDP-RaplB did not. By contrast, analysis of RaplA proteins in Epstein-Barr virus transformed human B lymphocytes found that both activating and dominant negative mutant proteins but not wild type Rap blocked the phorbol ester stimulated oxidative burst (Maly et al., 1994). This result may mean that in some cases the cycling of Rapl from a GTP to GDP bound form is required for activity.  Regulation of the Rapl proteins Like Ras, Rapl proteins are regulated by exchange factors and GAPs. A Rapl specific GAP (Rapl-GAP) that shows no sequence homology with RasGAP has been isolated and cloned from bovine brain (Rubinfeld et al., 1991). Rapl-Gap is phosphorylated  in vivo  and can be phosphorylated  in vitro  by  cAMP dependent protein kinase and also by p34 cdc2 (Polakis et al., 1992). The effect of phosphorylation on Rapl-GAP is not known. An exchange factor, smgGDS, which stimulates the rate of Rapl GDP/GTP exchange has been cloned (Kaibuchi et al., 1991). As mentioned previously, smgGDS is also active on other Ras related proteins (Kaibuchi et ah, 1991). To date, no specific downstream effectors of the Rapl proteins have been identified, although  RaplA has been shown to enhance protein kinase C activity in an in vitro assay (Labadia et al, 1993). Rapl also interacts with the Ras regulator Ras-GAP but Ras-GAP and NFI do not stimulate the GTPase activity of Rapl (Freeh et al, 1990; Zhang et al., 1991). In addition, Rapl interacts with the Ras effector, Raf, in a yeast two hybrid assay (Zhang et al, 1993). Whether such non-functional interactions are responsible for the antagonism of Ras transformation is not yet clear and it remains to be determined whether Rapl can interact with other recently identified Ras effectors or the Ras exchange factor. It is also plausible that Ras and Rapl act via the same effectors in cases where Rapl and Ras have similar effects. For example, human Rapl is capable of activating the Ras effector adenylate cyclase in S. cerevisiae (Ruggieri et al., 1992). Rapl is a substrate for protein kinase A and protein kinase G in vitro (Kawata et al, 1989; Miura et al, 1992). Phosphorylation of RaplB occurs in platelets upon treatment with agents such as prostaglandins which increase intracellular cAMP, suggesting that protein kinase A may phosphorylate Rapl in vivo (Kawata et al, 1989; Siess et al, 1990). The smg GDS induced activation of RaplB is enhanced upon phosphorylation of RaplB in vitro (Itoh et al, 1991), suggesting a mechanism whereby phosphorylation may regulate Rapl.  Summary Given the diverse effects of Rapl proteins and their sometimes antagonistic, sometimes complementary relationship with Ras and the failure thus far to detect a definitive Rapl specific effector, it remains to be determined if Rapl proteins function in the Ras signal transduction pathway by using or sequestering some of the components of that pathway or if Rap  acts in an independent pathway. It is unclear whether the diverse effects of Rapl proteins reflects a diversity of molecular interactions, or whether a single role for Rapl underlies these numerous effects.  A c t i o n of Ras superfamily proteins o n cell m o r p h o l o g y  The cytoskeleton of a cell is required for the structural integrity and shape of the cell and consists of a dense network made up of numerous components. These cytoskeletal elements form a matrix with a measurably rigid structure capable of resisting mechanical stress in a manner analogous to the rigid strength of geodesic domes designed by R. Buckminister Fuller (Wang et al, 1993; Heidemann, 1993). This matrix must also by necessity be capable of rapid disassembly and reassembly as a cell undergoes processes such as cellular division, developmentally regulated changes in shape, cell movement, and chemotaxis. Dramatic cytoskeletal changes also accompany the process of cellular transformation and occur during the treatment of cells with growth factors. Electron microscopy analysis has revealed that the cytoskeleton consists of 3 primary structures: microtubules, intermediate filaments and microfilaments.  Biochemical analysis has shown that  microtubules consist predominantly of tubulin; intermediate filaments are made up of a mixture of proteins; and microfilaments are made up of actin. I will restrict my focus to the actin cytoskeleton because it is subject to modification by Ras-related proteins. The  actin cytoskeleton consists of monomeric actin subunits  assembled into fibers which are in turn linked to create a matrix by actin crosslinking proteins (reviewed in Matsudaira, 1991). Many other actin associated proteins regulate the degree of actin polymerization: by preventing  the addition of further subunits to the fiber; by sequestering free actin subunits so as to prevent their assembly; and by severing actin fibers (Luna and Condeelis, 1990; Button et al, 1995). Actin filaments are present in numerous distinct forms in different subcellular structures in mammalian cells (reviewed in Matsudaira, 1991). Stress fibers consist of long bundles of actin filaments that traverse the cell and are linked to the extracellular matrix via integrins and focal adhesion complexes. Focal adhesions are involved in cell-substratum adhesion and consist of protein complexes linked to stress fibers. Lamellipodia are broad but thinly spread actin containing regions observed at the periphery of adherent cells. Membrane ruffles are similar to lamellipodia and are made up of actin containing wavy curtain-like structures on the cell surface and cell periphery.  Filopodia are narrow peripheral structures containing  'microspikes' of polymerized actin and are proposed to have a sensory function. As well as regulation by the assembly, disassembly and crosshnking of actin, the actin cytoskeleton is acted upon by force-generating myosin molecules. The conventional force-generating myosin, myosin II, forms filaments which bind to actin fibers and generate contractile forces (reviewed in Spudich, 1994; Ruppel and Spudich, 1995). Myosin molecules are involved in processes including muscle-driven movement, maintaining cell morphology and cell motility. In D.  discoideum,  myosin II is required for a  normal cell morphology (Knecht and Loomis, 1987; De Lozanne and Spudich, 1987). Many unconventional or nonfilamentous myosins have also been isolated (classes I, and III-VIII) (Titus et al, 1994; Bement et al, 1994). The roles of such unconventional myosins have not been determined to date. It has been estimated that a typical mammalian cell has at least 11 myosins  consisting of 1 or 2 members from each myosin class and it has been speculated that each myosin family member may mediate a different actinbased process (Bement et al, 1994).  The effects of the Ras superfamily on the cytoskeleton Structures such as membrane ruffles, stress fibers and focal adhesions are produced in response to growth factors and appear to involve members of both the Ras and Rho sub-families. Microinjection of activated H-Ras into fibroblasts results in a refractile transformed morphology and increased membrane ruffling and blebbing (Feramisco et al., 1984; Bar-Sagi and Feramisco, 1986). Microinjection of activated K-Ras into Swiss 3T3 cells also causes an increase in membrane ruffling and a decrease in stress fiber formation (Yoshida et al., 1992). Interestingly, microinjection of RaplB bound to GTPyS also caused membrane ruffling in Swiss 3T3 cells but did not have a significant effect on stress fiber formation. The relationship between Rapl and Ras in causing these partially similar phenotypes is not known. Microinjection studies have also identified roles for a number of Rho sub-family members in regulation of the cytoskeleton. Microinjection of an activated RhoA protein into serum starved Swiss 3T3 cells causes rapid stress fiber and focal adhesion formation (Ridley and Hall, 1992). Addition of either serum or lysophosphatidic acid caused a similar effect, suggesting that RhoA may mediate some of the effects of growth factors on the cytoskeleton. Specifically blocking RhoA function in  vivo  by microinjection of a dominant  negative form of RhoA, generated by ribosylation with the exoenzyme C3 transferase from Clostridium  botulinum,  or C3 transferase alone blocked the  effects of serum or lysophosphatidic acid, suggesting that RhoA is essential for  the assembly of focal adhesions and stress fibers induced by growth factors (Ridley and Hall, 1992). Microinjection of an activated Racl into Swiss 3T3 cells stimulates actin filament accumulation at the plasma membrane, forming lamellipodia and membrane ruffles (Ridley et al, 1992). Induction of membrane ruffling by growth factors or an activated H-ras protein could be blocked by a dominant inhibitory mutant Racl protein, suggesting that endogenous Racl is required for growth factor induced membrane ruffling. In addition, a delayed response to Racl is the formation of actin stress fibers. The results of blocking Rho activity with C3 transferase suggested that growth factors act through Racl to stimulate a Rho dependent response (stress fiber formation) as well as a Rho independent response (ruffling). Analysis of the human Cdc42 protein revealed that it too can regulate actin structures. Microinjection of human Cdc42 protein into Swiss 3T3 cells promoted the formation of peripheral actin microspikes and filopodia (Kozma et al, 1995; Nobes and Hall, 1995). Treatment of cells with the growth factor bradykinin had a similar effect which was blocked by microinjection of a dominant negative form of Cdc42. This result suggested that the endogenous Cdc42 protein is required for the formation of these peripheral actin structures. Analysis of the cells microinjected with Cdc42 also caused the formation of ruffles as well as stress fiber formation suggesting that perhaps Rac and RhoA were also being activated. Inactivation of the endogenous Rac and Rho proteins with a dominant negative Racl protein and C3 toxin blocked the formation of these structures, suggesting that Cdc42 acts in part via Rac and Rho. A model summarizing the relationship between the three Ras-related proteins is shown in Figure 2 (Chant and Stowers, 1995). The model is based  on the two lines of evidence discussed above. Firstly, the pathway appears to be branched because upstream components such as Cdc42 cause multiple effects while downstream components such as Rho cause a single effect. Secondly, the proteins appear to function in a linked manner rather than independently, because dominant negative forms of the presumptive downstream proteins such as Rho block some but not all of the effects of upstream proteins such as Cdc42. An additional level of complexity to the model is that Rac and Rho can also be activated independently by growth factors, but this can be explained by the branched nature of the pathway. A precedent for such a complex model linking multiple Ras-related proteins comes from the genetic analysis of bud site selection in S. (Fig. 2) (reviewed in Chant and Stowers, 1995). In S.  cei'evisiae,  cerevisiae  the site of the  daughter bud is not randomly selected but depends on the GTPase cycle of the Rapl homologue, RSR1/BUD1, which is linked in turn to the GTPase cycle of Cdc42. A series of Rho proteins are then proposed to function downstream of Cdc42 and perhaps act on the cytoskeleton itself. The protein Cdc24 potentially links RSR1/BUD1 and Cdc42, because it binds to RSR1/BUD1 and also functions as an exchange factor for Cdc42.  Figure 2. GTP cascades controlling cell morphology and bud formation This figure is modified from (Chant and Stowers, 1995). (A) The pathway that controls cytoskeletal polarization during vegetative division or mating. (B) The proposed pathway that controls the formation of filopodia (microspikes), lamellipodia (ruffles) and stress fibers with focal adhesions. Continuous arrows indicate physical contact. Broken arrows indicate indirect interactions.  ^  Intrinsic spatial signals for budding  Mating pheromone  PDGF or  B  LPA  insulin  RAS  t BUD5 (CDC42) (  BUD1  BUD2  XT  V  Rlopodia  CDC24 GEF  C RAC  CCDC42)  Lamellipodia C RHCy RH01 RH02 RH03 RH04  Cytoskeletal polarization  Stress fibers Adhesion  Dictuostelium  discoideum  Life cycle D . discoideum  is a simple eukaryote which serves as an experimentally  tractable model system for addressing many biological questions. D . discoideum  cells exist as solitary amebae that feed on bacteria but, when  starved, aggregate and form a developmental structure (reviewed in Gross, 1994; Kay, 1994; Firtel, 1995) (Fig. 3). The cells cease growing when deprived of nutrients and signal to each other via pulses of cAMP. Cells respond by moving towards the stimulus and by producing cAMP themselves. This chemotaxis and signal relay results in the formation of an aggregate of cells. The aggregate becomes compact and tipped and the tip eventually elongates to form a finger-like structure. This structure falls over and continues to elongate producing a pseudoplasmodium or slug. The slug is capable of migrating towards conditions of optimal temperature and light. The slug tip consists of prestalk cells which make up approximately 20% of the cell total and the rear 80% of the slug consists of prespore cells. The ratio of prestalk to prespore cells is fixed. If the front or back of the slug is removed experimentally, the remaining cells will again assume the proportions of 20% prestalk 80% prespore cells. The maturation of the structure occurs as the prestalk cells at the tip of the structure migrate down through the center of the prespore cell mass in a motion which resembles that of an inverse fountain. The consequences of this prestalk cell movement is the formation of a stalk which raises the prespore mass away from the substratum. During this process the prespore cells develop into mature spores.  Figure 3. The life cycle of D.  discoideum  Black areas represent prestalk and stalk cells while clear areas represent prespore and spore cells. The time in development is shown below in hours.  S i g n a l transduction events d u r i n g the lifecvcle of D.  discoideum  Vegetative growth, starvation and aggregation Vegetative  D. discoideum  cells prey on bacteria by detecting and  chemotaxing toward folate and pterin, two bacterial metabolites (Pan et al, 1972; Pan et  al,  1975). The mechanism that D.  cells use to  discoideum  respond to folate and pterin is not well understood, although it is known to require the G(3 and Ga4 members of the heterotrimeric G protein family (Wu et al, 1995; Hadwiger et al, 1994; Burdine and Clarke, 1995). Folate binding sites have also been detected on the surface of D. 1982; De Wit  etal,  discoideum  cells (De Wit,  1985).  To determine when to stop growing and initiate  collective  development, cells evaluate both their nutritional status and also their density. This latter process is achieved by secreting and responding to a prestarvation factor (PSF) (Clarke et al, 1987; Clarke et al, 1988). Another factor, conditioned medium factor (CMF) provides a similar density sensing function which regulates signal transduction after starvation has initiated (Yuen et al., 1995). After cells starve and enter into development, the reintroduction of nutrients will cause the cells to dedifferentiate although such cells retain the capacity to rapidly reenter the developmental program if nutrients are removed for a second time (Waddell and Soil, 1977; Soli and Waddell, 1975; Kraft et al, 1989). This 'memory' exists for approximately 90 minutes and is then lost by a process that has been termed erasure. Presumably, such flexibility reflects selection pressures of living in an environment where the food supply is variable and intermittent. The cAMP signal relay response mediating the process of aggregation and early gene expression has been extensively analyzed (reviewed in Gross,  1994; Firtel, 1995). The signal relay machinery consists of a serpentine cAMP receptor, cARl, linked to a heterotrimeric G protein made up of Ga2, Gp and presumably a hitherto unidentified Gy subunit. Signal transduction also appears to involve the protein kinase ERK2. Starving cells emit and respond to pulses of cAMP. The binding of cAMP to receptors activates phospholipase C, guanylate cyclase and Ca2+ influx. Protein kinase A, presumably activated by increased intracellular cAMP, has also been shown to be required for aggregation. Cells chemotax toward cAMP and in addition respond by activating adenylate cyclase, which produces further cAMP allowing a 'relay' of the signal. Cells also secrete a cAMP phosphodiesterase which degrades extracellular cAMP, preventing its over-accumulation. Cells also respond to cAMP signals by expressing early developmental genes.  Cell differentiation The differentiation of cells into prestalk and prespore cells within the multicellular aggregate is regulated by a number of factors including cAMP (reviewed in Gross, 1994; Firtel, 1995). Levels of cAMP increase during development and the presence of continuously high levels of cAMP represses the expression of genes involved in early development and stimulates expression of genes involved in later processes. cAMP initially promotes the development of both prestalk and prespore cells but at later stages acts to promote spore development and inhibit stalk development.  Four  developmentally regulated cAMP receptors and 8 heterotrimeric Ga protein subunits have been identified, suggesting the potential for several distinct cAMP pathways. Protein kinase A is required for normal development and has been shown to be a positive regulator of prespore gene expression and spore maturation, presumably functioning to mediate the effects of  intracellular cAMP (Simon et al, 1989; Firtel and Chapman, 1990; Harwood et al, 1992a; Harwood et al, 1992b; Simon et al, 1992; Anjard et al, 1992; Mann et al, 1994; Hopper et al, 1995). Protein kinase A also appears to inhibit prestalk development and to play a role in stalk formation (Mann and Firtel, 1993; Harwood et al, 1992b). An additional factor, stalk cell differentiation inducing factor (DIF) promotes prestalk cell differentiation and suppresses prespore cell differentiation (Kopachik et al., 1983; Williams et al., 1987). Finally, ammonia, which accumulates as cells catabolize protein and RNA by endogenous respiration during development, serves as a negative regulator for the process of culmination - the formation of a mature fruiting body from a slug (reviewed in Gross, 1994). Although the factors which promote the development prestalk and prespore cells have been identified, the question of how the proportions of the two cell populations is determined remains unanswered. Prestalk and prespore cells have been observed to start differentiating in an interspersed manner within the early aggregate, suggesting an initially cell-autonomous process rather than a response to an external gradient (Williams et al, 1989). Subsequently there is a migration of prestalk cells to the tip of the aggregate and this is believed to be in response to cAMP (Traynor et al, 1992). Within the aggregate and slug, exposure to morphogens promotes the continued development of prestalk and prespore cells. At least three prestalk cell types, as defined by differential patterns of gene expression, have been identified (Williams et al., 1989; Jermyn et al., 1989). There are also interactions between the prestalk and prespore cells which result in the regulation of the final proportion of the two cell populations (Raper, 1940; Shaulsky and Loomis, 1993). Factors such as position within the cell cycle during vegetative growth prior to the initiation of development also influence cell fate (Maeda et al.,  1989; Gomer and Firtel, 1987). A number of models have been proposed to explain the proportioning of the stalk and spore populations, utilizing some or all of the morphogens described above and invoking gradients of the morphogens (reviewed in Gross, 1994). However, to date, evidence for the spatial distribution of the above morphogens is not convincing (Weeks and Gross, 1991).  Ras a n d ras related genes i n D.  D. discoideum et ah,  1989; Daniel et  discoideum  expresses five al.,  ras  genes (Reymond et  1993b; Daniel et al, 1993a).  rasG  al.,  1984; Robbins  encodes a protein  which shares 69% overall amino acid identity with the human H-ras gene product, and the gene is expressed in vegetative and early developing cells with expression declining markedly during aggregation (Robbins et al., 1989). The RasB and RasC gene products share 59% and 56% amino acid identity, respectively, with the human H-ras gene product (Daniel et al., 1993a; Daniel et al,  1993b).  rasB  is maximally expressed during vegetative growth and early  development but expression remains relatively high during the remainder of development (Daniel et  al,  1993b).  rasC  is expressed maximally during  aggregation but significant expression is detected during vegetative growth and the remainder of development (Daniel et al, 1993a). Two additional Ras genes, rasS and rasD, whose gene products share 54% and 65% amino acid identity, respectively, with H-ras, are expressed in a more restricted manner (Daniel et  al,  1993a; Reymond et  aggregation (4-8 hours) while  al,  rasD  1984).  RasS  is expressed only during  is highly expressed only during late  aggregation and slug formation (12-16 hours).  rasD  gene expression is  induced by cAMP and is expressed in at least 50% of cells during aggregation  but is subsequently restricted to the prestalk cells by an unknown mechanism (Jermyn and Williams, 1995). The existence of 5  ras  genes in D.  discoideum  each with a specific pattern of expression, raises the question of whether they each have distinct roles and this question remains to be answered. The presence of amino acid differences within the core effector domain of RasC and RasS may mean that they interact with different effectors than do the other three Ras gene products (Daniel et al, 1993a). The Ras proteins have not been definitively linked to any specific growth or developmental signaling pathway in D. discoideum.  Expression of  an activated RasG protein in vegetative cells under the control of the discoidin  promoter blocks aggregation and northern blot analysis showed  only low levels of induction of the cAMP receptor, cARl, which is normally expressed early after only 2-4 hours development, suggesting a role in early development (M. Khosla, personal communication). Expression of an activated rasD gene under the control of the rasD promoter resulted in cells which were arrested as multitipped aggregates (Reymond etal,  1986).  Northern blot analysis showed a dramatic increase in prestalk specific  ecmA  mRNA levels and a decrease in prespore specific sp60 mRNA levels (S. Louis, personal communication), indicating that expression of activated  rasD  disrupts pattern formation. Eight rho and 5 rab genes have been isolated from D.  discoideum  (Bush  et al, 1993a; Bush et al, 1993b; Vithalani et al, 1995). The rho related genes each exhibit unique patterns of mRNA expression during growth and development, again consistent with the idea that they may have different roles during the D.  discoideum  life cycle. In contrast, four of the five  rab  genes have similar expression profiles. Disruption of one rho gene interfered  with cytokinesis (Vithalani et al, 1995) but the roles of the remaining rho and rab genes remain to be established. Only one rap gene, rapl, has been identified in D. discoideum (Robbins etal.,  1990) despite extensive searching for additional  rapl  genes (Daniel,  1993). The Rapl gene product shares 76% amino acid identity with the human Rapl A protein and approximately 50% amino acid identity with the H-ras gene product (Robbins et al., 1990; Daniel, 1993). A complex pattern of rapl  mRNA expression is observed (Robbins et  al.,  1990). A single 1.1 kb  mRNA is present during vegetative growth and early development. As development continues, this message is replaced by two mRNAs of 1.0 and 1.3 kb. mRNA levels are maximal during aggregation, then decline during slug formation, but increase again slightly at culmination. Separation of prestalk and prespore cells by a percoll gradient followed by mRNA analysis did not show any enrichment in either prestalk or prespore cells (Robbins, 1991). However, in contrast to the complex developmental pattern of mRNA expression, western blot analysis showed that protein levels remained constant throughout growth and development (M. Khosla, personal communication).  Cell morphology of D.  D.  discoideum  discoideum  has become an important experimental model for  analysis of the function of actin associated proteins and myosin motors, using techniques for gene targeting and gene replacement (Patterson etal, 1991). The actin-associated proteins a-actinin, ABP-120, coronin, synexin, ponticulin and profilin have each been eliminated by gene disruption techniques (Witke et al, 1987; Cox et al, 1992; de Hostos et al, 1993; Hitt et al, 1994; Doring et al,  1991). Similarly, several myosin I genes have been disrupted (Jung et al, 1993; Jung and Hammer III, 1990; Peterson et ai, 1995). Many of these gene disruption mutants strains appear morphologically normal and exhibit only subtle changes in their motility. However, disruption of the myosin II light or heavy chain genes results in cells which are enlarged, flattened and multinucleate (Knecht and Loomis, 1987; De Lozanne and Spudich, 1987). Such cells are incapable of division in suspension but continue to grow until they lyse. These cells are no longer able to cap surface proteins, have reduced cortical tension and no longer respond to azide treatment by contracting and detaching from the substratum (Pasternak et al, 1989). Surprisingly, such cells are still motile and can stream, aggregate and develop to the mound stage, although development arrests at this stage (Knecht and Loomis, 1987; De Lozanne and Spudich, 1987; Springer et al, 1994). This range of defects suggests that myosin II is involved in cytokinesis, receptor capping, control of cell morphology and morphogenesis. Some of the components involved in regulating myosin II in D. discoideum  have been identified. Myosin II is structurally and functionally  similar to non muscle myosin II in other organisms (reviewed in Tan et al., 1992; Spudich, 1994). Myosin II consists of a hexamer of subunits (2 heavy chains, 2 essential light chains and 2 regulatory light chains) forming a structure with two globular heads and a helical coiled-coil tail. The two globular heads consist of the amino terminals of two heavy chains and the myosin essential light and regulatory light chains which the tail consists of the carboxyl terminals of heavy chains. The hexamers assemble into filaments which are capable of contracting, and which are analogous to those in muscle.  Both myosin heavy chain and regulatory light chains are subject to phosphorylation (reviewed in Tan et al., 1992; Hammer III, 1994). Myosin heavy chain phosphorylation occurs on multiple residues in the tail and this prevents assembly of filaments capable of exerting force. Amino acid substitutions which mimic the phosphorylated state of the myosin II heavy chain tail prevent the assembly of myosin onto the actin cytoskeleton, while disruption of three sites of phosphorylation results in the overassembly of myosin II on the actin cytoskeleton  in vivo  (Egelhoff  et al,  1993). To date,  three myosin heavy chain kinases have been identified and one of these is a member of the protein kinase C family (Maruta et al., 1983; Cote and Bukiejko, 1987; Ravid and Spudich, 1989; Ravid and Spudich, 1992). Myosin light chain is phosphorylated by myosin light chain kinase which causes an increase in actin activated ATPase activity of myosin II in  vitro  (Griffith et  al.,  1987). Phosphorylation of myosin light chain does not regulate the assembly/disassembly of myosin II, a situation different from that in higher eukaryotes (Tan et al, 1992). As  D. discoideum  amebae start to chemotax toward cAMP, they become  much more elongated, and while streaming together make head to tail contacts. Observation of single cells responding to a pulse of cAMP has revealed that cells initially cringe before extending a pseudopod in the direction of the cAMP stimulus (Futrelle et al., 1982). There is a rapid accumulation of F-actin into the cytoskeleton (peaking 5 s after the cAMP stimulus) and phosphorylation of the myosin II heavy chain accompanied by an association of myosin II with the cytoskeleton, occurs 25-30 s after the cAMP stimulus (McRobbie and Newell, 1984; McRobbie and Newell, 1983; Berlot et al., 1985; Berlot et al., 1987). The myosin light chain is also phosphorylated following cAMP treatment (Berlot et al., 1985). The transient  increase in intracellular cGMP which occurs following cAMP stimulus has been shown to be important for the regulation of the myosin II responses (Liu and Newell, 1991; Liu etal., 1993; Liu and Newell, 1994). Strains with a defective cGMP-specific phosphodiesterase have persistent elevated levels of cGMP following a cAMP stimulus and in these strains, there is prolonged association of myosin LI with the cytoskeleton and a delay in myosin II light and heavy chain phosphorylation. It has been proposed that guanylate cyclase is activated via Ca2+ influx (Newell and Liu, 1992). However, the pathways mediating the phosphorylation of myosin via myosin kinases in response to cAMP have not been fully elucidated. The response of starved cells to nutrients has some similarities to the cAMP cringe response; When nutrient medium is reintroduced to starved cells, the ameboid cells rapidly round up and become refractile in a process which has been compared to the response of mammalian cells to serum (Schweiger et al, 1992; Howard et al., 1993). The HL5 stimulation response is accompanied by a rapid tyrosine phosphorylation of a subset of cellular proteins including actin (Schweiger et al, 1992; Howard et al, 1993). Disruption of tyrosine phosphatase PTP1, but not PTP2, caused a more rapid and more prolonged phosphorylation of actin and an acceleration of cell rounding when starved cells were returned to growth medium (Howard et al., 1993; Howard et al., 1994). Overexpression of PTP1 decreased the amplitude and duration of actin phosphorylation and also diminished the cell rounding response (Howard et ai, 1993). In addition, treatment of vegetative cells with the phosphatase inhibitor, phenylarsine oxide, caused cell rounding accompanied by actin phosphorylation (Schweiger et al, 1992). Although some of the components involved in the cAMP signal transduction relay in D .  discoideum  have been identified, the details of the  signal transduction to the cytoskeleton are not known. Similarly the response of starved cells to nutrients has not been well characterized. One strategy for studying the control of the cytoskeleton is to identify signal transduction molecules which, when disrupted or overexpressed, affect cell morphology and motility. Overexpression of Gal or the expression of a calmodulin antisense RNA both produce large multinucleate cells that are reminiscent of those caused by disruption of myosin II heavy chain and profilin (Kumagai et al, 1991; Liu et al, 1992).  Rationale and research objective At the outset of this work, the function of the D. discoideum  rapl  gene  was unknown. The objective of my research has been to further characterize the organization of the rapl gene and to evaluate its role in D. discoideum by overexpression of wild type and mutant Rapl proteins.  Site-directed  mutations were generated in rapl to target conserved amino acids previously shown to be important for activity in either mammalian Rapl or Ras proteins or both. During the course of this work, I noted that a transformant expressing an activated RasG-G12T protein (generated by M. Khosla) shared some of the characteristics of transformants overexpressing the Rapl protein, and therefore I characterized this transformant in more detail.  MATERIALS AND METHODS Materials G418 (Geneticin) was purchased from Sigma (USA), FITC-phalloidin was purchased from Molecular Probes (USA) and X-ray film was purchased from Kodak (Canada). Radiolabeled [oc32p] dCTP was purchased from ICN How Labs (Canada) and [35s] dATP was purchased from Dupont NEN Canada Inc., filters were purchased from Millipore (USA) and the enhanced chemiluminescence kit for western blot analysis was purchased from Amersham (Canada). Bacteriological peptone and yeast extract were purchased from Oxoid (UK). All other chemicals were purchased from Fisher ScientiEc Co. (USA) or BDH (Canada). Hoechst 33258 dye was a gift from Dr. Hancock's laboratory. Oligonucleotides were synthesized by Dr. Sadowski's laboratory (UBC) or the NAPS unit (UBC). Restriction endonucleases or modifying enzymes were purchased from GIBCO BRL (Canada). Taq polymerase was purchased from Promega (USA); Vent polymerase was purchased from New England Biolabs (USA) and Sequenase was purchased from United States Biochemical. The anti-Rapl peptide antibody and the anti-RasG-GST-fusion protein antibody were generated by Steve Robbins (Robbins, 1991). The antiphosphotyrosine antibody IgG2bk was initially purchased from Upstate Biotechnology Inc. (USA) but subsequently was a gift from Dr. Mike Gold. Goat anti-mouse IgG antibody conjugated to horseradish peroxidase and goat anti-rabbit IgG antibody conjugated to horseradish peroxidase, the secondary antibodies for ECL analysis, were purchased from Amersham (USA). The  E. coli  strain DH5ctF' was used for bacterial transformations. The  genotype of DH5aF' is:  F/endAl  //sdRi7(rk"mk ) +  supE44  thi-1  iecAl  gyrA(Nair) relAl  A(lacZYA-argF)U169  deoR (08OdlacA(lacZ)Ml5)  (Raleigh et  al, 1989). The pVEH vector was donated by Wolfgang Nellen, the pDdGal 17 vector was donated by Adrian Harwood and the pVEIIGal vector was donated by Birgitte Wetterauer.  D. discoideum  Growth of D.  growth and differentiation  discoideum  The Ax2 strain of D.  discoideum  that was used in all experiments was  grown axenically in HL5 medium (Watts and Ashworth, 1970) (14.3 g neutralized bacteriological peptone, 7.15 g yeast extract, 0.96 g Na2HPC»4 and 0.486 g KH2PO4 per liter of water) with gyratory shaking at 150 rpm at 22°C or on plates in association with E. aerogenes. Cell numbers were determined in duplicate with a hemacytometer.  The transformed Ax2 strains were  maintained in HL5 medium in the presence of 10 /ig/ ml G418 (Geneticin) except the wild type Rapl transformant (containing the pVEII Rapl vector) (Rebstein et al, 1993) which was maintained in the presence of 50 ug/ ml G418.  Development of D. D.  discoideum  discoideum  development on filters was initiated as previously  described (Khosla et al, 1990). Exponentially growing cells were washed twice in BS buffer (10 mM NaCl, 10 mM KC1 and 2 mM CaCl2) (Bonner, 1947) by centrifugation at 700g for 3 minutes and 2.5 X 10^ cells were plated on a 4.0 cm diameter filter (pore size = 0.45 um), resting on a BS buffer-saturated pad in a 60 mm petri dish. The filters were incubated at 22°C in a moist chamber. In some experiments, cells were developed in suspension in an Erlenmeyer flask at a density of 2 X10^ cells/ml with gyratory shaking at 150 rpm at 22°C. D.  discoideum  development following growth on bacteria was  accomplished by pipetting 1-5/d of cells in HL5 medium onto a freshly inoculated lawn of Enterobacter  aerogenes  on an SM nutrient agar plate (10 g  glucose, 10 g neutralized bacteriological peptone, 1 g yeast extract, 1 g  MgS04-7H20,1.55 g NaH2P04 H20,1 g K H P O 4 and 20 g bactoagar per liter of water). Plates were incubated at 22°C and after the D.  discoideum  cells had  consumed the bacteria (usually 4 days), development ensued in the zone depleted of bacteria. To analyze the erasure response, a modification of a previously described procedure was used (Kraft et al., 1989). Cells were starved in BS buffer in suspension, as described above, for 8 hours, incubated in HL5 medium for either 1 or 2 hours under shake condition, washed twice in BS buffer and then allowed to develop on filters as described above. Determination of cell viability Cells were grown to high density in the absence of folate, washed twice, and approximately 100 cells were plated in 100 mm tissue culture plates. After allowing the cells to adhere for 30 minutes, HL5 was added back either immediately or after 8 hours of starvation. Cells were counted after 6 days. The percentage of cells viable after starvation was determined by counting the number of CFU before and after starvation on duplicate plates.  Induction of the  discoidin  promoter  To maintain strains containing genes under the control of the discoidin  dis  I y gene promoter in a suppressed state, 1 mM folate was added  to the HL5 medium. To induce expression from the discoidin  promoter, cells  were incubated with conditioned HL5 medium, since conditioned medium contains a pre-starvation factor (PSF) (Clarke et al, 1987; Clarke et al, 1988), that induces expression from the  discoidin  promoter (Rathi et  al,  1991).  Conditioned HL5 medium was prepared by growing Ax2 cells to a density of  approximately 2 x 10^ cells/ml, removing the cells by centrifugation and filtering the medium though a 0.2 um pore size nitrocellulose filter. The  discoidin  promoter is also induced between 4 and 8 hours of  development (Blusch et al, 1992), which allows the effects of genes expressed from this promoter to be analyzed during early development. To ensure continuous high expression of the Rapl and RasG proteins from the start of development, cells were grown under inducing conditions in conditioned medium for 24 hours prior to the initiation of development.  In some  experiments, induction was achieved by growing cells to 1-3 xlO^ cells/ml in the absence of folate. Development was then initiated as described above.  Analysis of cell morphology To examine cell morphology, cells were plated at 3x10^ cells/ cm^ on a glass coverslip in a 60 mm petri dish and incubated for 24 hours either in the presence of 5 ml of conditioned HL5 medium or in HL5 medium containing 1 mM folate to induce or repress the  discoidin  promoter, respectively. To  evaluate the ability of the cells to undergo the nutrient stimulus induced cell rounding, the cells were then starved for 8 hours in 5 ml BS buffer and then exposed to 5 ml of HL5 medium. Cells were photographed before HL5 stimulation and then 5 and 10 minutes afterwards. To determine the effect of azide on cell substratum adherence, an adherent monolayer of cells was exposed to 2 mM Na azide (NaN3) in HL5 medium for 3 minutes with swirling at 60 rpm, in a modification of a previously described procedure (Pasternak et al, 1989; Springer et al, 1994). The plates were gently rinsed 3 times to remove floating cells. The coverslip was photographed before and after azide treatment. Between 200 to 600 cells were counted prior to azide treatment.  Analysis of cell-cell adhesion Cell adhesion was measured using an agglutometer, which monitors light scattering (high values indicate small aggregates or single cells while small numbers indicate large aggregates). Cells were grown to a density of 2xl06 cells/ml, washed in PBS and assayed for cell agglutination over a 60 minute period in the presence and absence of cell adhesion.  Flow cytometric analysis Cells were grown in conditioned medium for 24 hours in shake suspension and analyzed directly in medium using a flow cytometer (BectonDickinson) running Lysis II software. Five thousand events were analyzed for forward and side light scatter for each sample. Mean values were determined for forward scatter events above a cutoff point of 200.  Transformation of D. discoideum  D.  discoideum  Ax2 cells were transformed by the calcium phosphate  precipitation technique (Nellen et al., 1984) in Bis-Tris HL5 (Egelhoff et al., 1989). After incubating the calcium phosphate DNA precipitate with the cells for 4 hours, the cells were given a 2 minute osmotic shock with 15% glycerol as previously described (Early and Williams, 1987). Transformants were selected in 10/tg/ ml G418 in HL5 and colonies were visible after approximately 10 days. Individual clones were transferred initially to 24 well plates, then to 100 mm plates. Established transformants were maintained in shake suspension as described above. The RasG,  RasG-G12T,  and  RasG-SUN  transformants were generated in a similar manner (Khosla et al., 1995).  Dark field, Nomarski andfluorescencemicroscopy Vegetative cell morphology, the nutrient-induced rounding of starved cells and the effects of azide treatment were observed by darkfieldmicroscopy of an adherent layer of living cells on a glass coverslip. A two-sample t test for the two tailed hypothesis was used to test the significance of differences observed in the means between selected samples. Relative cell areas were determined by digitizing photographs of the cells (pdi Model DNA 35 scanner), manually tracing the outline of the cells on the digitized image and computing the areas with the PD Quest 4.1 program. To observe F-actin distribution, cells adherent to a glass coverslip were fixed with 3.7% formaldehyde in BS buffer for 10 minutes and then washed 3 rimes with PBS (8 g NaCl, 0.2 g KC1,1.44 g NaHP04,0.24 g K H 2 P O 4 in 1L H 2 O pH 7.4). Cells were then permeabilized for 5 minutes in -20°C acetone, rehydrated in PBS, overlaid with FITC-phalloidin solution (0.2 yM) and then washed with PBS to stain F-actin. To stain cell nuclei, the formaldehyde fixed cells were overlaid with 0.0005% Hoechst dye # 33258 for 5 minutes and then washed with PBS (Harlow and Lane, 1988). Cells were viewed using a Zeiss Axiophot microscope equipped with epifluorescence. All photographs were taken using TMAX 400 film.  Molecular Biology  Plasmid DNA preparation Plasmid DNA was isolated from DH5cxF'  E. coli  cells using a CTAB  miniprep procedure (Del Sal et al., 1989) or a PEG-precipitation large scale procedure (Maniatis et  al,  1989). Competent  D H 5 c t F ' E. coli  cells were  prepared using a rubidium chloride technique and transformations of  competent cells were done as described (Maniatis et al, 1989). Transformed DH5cxF E. coli cells were selected on LB ampiciUin plates (10 g bacto-tryptone, 5 g bacto yeast extract, 10 g NaCl, 15 g bacto-agar pH 7.0, with 60 Mg/ml ampicillin).  Southern blot analysis Genomic DNA was prepared as described (Maniatis et al, 1989) from isolated D. discoideum nuclei (Coccucci and Sussman, 1970). Briefly, isolated nuclei from 3.7 x 10^ cells were lysed in 1% SDS, treated with proteinase K (200/ig/ml), and extracted three times with an equal volume of phenolchloroform. The sample was then ethanol precipitated, resuspended and treated with DNase-free RNase (10/ig/ ml). The sample was extracted with an equal volume of phenol-chloroform, ethanol precipitated and then resuspended in TE (10 mM Tris HCl 1 mM EDTA pH 7.4). The genomic DNA concentration was determined spectrophotometrically (Maniatis et al, 1989). Six Mg of genomic DNA were digested overnight with 25 units of restriction enzyme and the DNA was separated on a 0.8% agarose TBE gel (IX TBE is 89 mM Tris-borate, 89 mM boric acid 2 mM EDTA). The DNA was stained with ethidium bromide and the gel was photographed. The DNA was then denatured in 1.5 M NaCl, 0.5 M NaOH for 30 minutes and then neutralized in 1.5 M NaCl, 0.5 M Tris pH 8.0 for 30 minutes. The gel was soaked in 2X SSC and then transferred and fixed to nitrocellulose. Nitrocellulose filters were hybridized overnight at 37°C with the rapl cDNA probe in hybridization buffer (5X SSC (IX SSC is 150 mM NaCl, 15 mM Na citrate, pH 7.0), IX Denharts (0.1% ficoll, 0.1% polyvinylpyrrolidone, 0.1% bovine serum albumin), 50 mM NaPC»4, 0.5% SDS and 30% formamide) (IX  SSC is 0.15 M NaCl, 0.015 M Na citrate) and then washed twice at 65°C in 0.1X SSC and 0.1% SDS. Filters were then exposed to X-ray film at -70°C.  Preparation of cDNA probes The cDNA fragment was purified by gel electrophoresis in a 2% low melt agarose gel. An aliquot of the low melt agarose containing the cDNA fragment was used directly for radiolabeling by nick translation (Maniatis et al., 1989). Nick translation was performed as described (Feinberg and Vogelstein, 1983). The radiolabeled probe was separated from unincorporated nucleotides with a Sephadex G-25 spin column (Maniatis et al, 1989).  Sequencing Single stranded DNA was produced by infection with K07 M13 helper phage, followed by PEG precipitation, isolation on glass filters and elution into TE (Maniatis et ah, 1989). Double stranded DNA was sequenced directly from CTAB miniprep DNA. Sequencing reactions were performed according to the manufacturer's (United States Biochemical) protocol except that the Sequenase reaction buffer was added after DNA denaturation. DNA sequence was determined by the dideoxy chain termination method (Sanger et al., 1977) with modified T7 DNA polymerase.  Electrophoresis and immunoblotting SDS-PAGE and immunoblotting techniques were performed as described (Robbins, 1991). For western blot analysis of transformants expressing genes under the control of the  discoidin  promoter, cells were  inoculated at a density of 5 xlO^ cells/ml and grown for 24 hours in shake suspension in either conditioned medium or HL5 containing 1 mM folate.  Cells were lysed in 1% SDS and diluted in an equal volume of 2X loading dye (20% glycerol, 10% p-mercaptoethanol, 4.6% SDS, 125 mM Tris HCl pH 6.8). Protein concentration was determined by UV absorbance (Harlow and Lane, 1988). About 10-15 ug of protein were electrophoresed on a 12% SDSpolyacrylamide gel and transferred to a nitrocellulose filter. Prestained molecular weight markers (BioRad) were used to estimate protein sizes. The nitrocellulose blots were stained with Ponceau S (Harlow and Lane, 1988) to confirm that equal amounts of protein had been loaded and transferred in all lanes. The membrane was blocked at room temperature with IX TBS (8 g NaCl, 0.2 g KC1,3 g Tris HCl in 1 L H2O pH 7.4) 5% skim milk and 1% Tween20. The Rapl protein was detected with a specific anti-Rapl peptide antibody at a 1:2000 dilution in IX TBS containing 0.5% skim milk and 1% Tween-20 (Robbins, 1991). The RasG protein was detected with a specific anti-RasG-GST protein antibody at a 1:5000 dilution (Robbins, 1991). A goat anti-rabbit antibody conjugated to horseradish peroxidase was used as a secondary antibody to generate a signal by ECL which was recorded on X-ray film. Protein phosphotyrosine western blot analysis was performed similarly using the anti-phosphotyrosine monoclonal antibody IgG2bk at a dilution of 1:2000 in IX TBS, 5% BSA, 0.1% Tween-20, 0.5 mM Na3V04 and 0.2 mM Na2Mo04A goat anti-mouse IgG antibody conjugated to horseradish peroxidase was used as a secondary antibody.  fr-galactosidase assay Growing cells were washed twice and 1x10? cells were plated in a 100 mm tissue culture plate in 10 ml KK2 buffer (20 mM KHPO4, pH 6.0) to initiate development. At the appropriate stages of development, the cells were rinsed in fresh KK2 buffer and resuspended in 0.5 ml 0.1 M phosphate  buffer pH 7.0. Cell-free extracts were prepared by subjecting the cells to three cycles of freezing in a dry ice-ethanol bath and thawing at 37°C. The broken cell suspension was centrifuged (RCF = 13600) to remove insoluble debris and the supernatant fluid was assayed for (3-galactosidase activity using onitrophenyl-p-D-galactoside (ONPG) as the substrate (Miller, 1972; Dingermann et al., 1989; Knox et al., 1991). |3-galactosidase activities from samples were determined from duplicate reactions and averaged. Protein concentration was determined using the Bradford assay with BSA as a reference standard (Bradford, 1976).  Isolation of the rapl genomic region A sucrose gradient from 10% to 40% sucrose was prepared with a gradient-forming device. Genomic DNA (50 /ig) was digested overnight with 50 units of Xbal and 50 units of BglTL, ethanol precipitated, dissolved in 100fd of TE buffer and then centrifuged on this gradient at 34000 rpm (RCF = 150 000) in an SW-41 swinging bucket rotor for 20 hours at room temperature. 1 ml fractions were collected and size analyzed by gel electrophoresis. DNA from the appropriate fraction (2-8 kb) was isolated by diluting the sample 4 fold in TE followed by ethanol precipitation. This fraction was predicted to contain the 6.6 kb 5' fragment and 4.0 kb 3' fragment of the rapl gene. The size-fractionated DNA was cloned into  BamHI  Xbal  digested gel  purified Bluescript vector, and approximately 35000 CFU were screened for inserts containing rapl DNA. The colonies were transferred to nitrocellulose using standard techniques and the colony blots were probed with a radiolabelled cDNA consisting of the full length rapl cDNA prepared by nick translation as described for the Southern blot procedure (Maniatis et al,  1989).  Briefly, nitrocellulose filters were hybridized overnight at 37°C with the  probe in hybridization buffer and then washed twice at 50°C in 2X SSC and 0.1% SDS. Filters were exposed to X-ray film and 15 positive clones were identified. The positive clones were picked, diluted, plated and the resulting colonies were probed as described above. Two colonies were still positive after probing and the presence of the predicted insert was confirmed by restriction digestion and Southern blot analysis. As both cloned inserts consisted of the 3' fragment of the rapl gene, a modified strategy was therefore used to isolate the remaining 5' fragment of the rapl gene. The size-fractionated genomic DNA was further digested with Hindi,  reducing the size of the presumptive insert to 1.5 kb and the fragments  were cloned into rapl  BamHI  Hindi  digested Bluescript vector. An EcoRI  Bglll  cDNA fragment specific for the 5' region of the gene was used as a probe.  Approximately 60000 clones were screened as described above. Five positive clones were identified, picked, diluted, plated out and the resultant colonies were probed as described above. Only one colony remained positive after this probing, but it was shown to contain the predicted insert by restriction digestion and Southern blot analysis. A missing internal 78 bp Bglll  Bglll  fragment was isolated by PCR. The  presence of the predicted 600 bp fragment was confirmed by running 5ul of the PCR reaction on a 0.8% agarose TBE gel. The remaining 45/il of the PCR reaction was ethanol precipitated with ammonium acetate and resuspended in TE. The PCR product was digested with  Bglll,  extracted with phenol  chloroform, ethanol precipitated and then resuspended in TE. The PCR product was ligated into BamHI digested Bluescript vector overnight at room temperature. The ligation mixture was further digested with  BamHI  to  linearize Bluescript vector which did not contain insert DNA, heated at 65°C for 10 minutes and then transformed into competent DH5aF' E coli cells.  Sequence analysis of the rapl genomic region A 1.8 kb Acc\ fragment from the 3'  rapl  fragment was cloned into the  Bluescript vector in both orientations for sequence analysis. The 5'  rapl  fragment in Bluescript was removed by SstI Hindll digestion, was gel purified and cloned into SstI  Hindll  digested pTZ18U. Nested deletions (Maniatis et  ah, 1989) of the genomic fragments were generated with exonuclease III in both orientations using the 3'  rapl  complementary orientations of the 5'  fragment in Bluescript, and in rapl  fragment in Bluescript and  pTZ18U, respectively. Briefly, DNA was digested with two restriction enzymes to linearize the DNA and create a single end resistant to exonuclease IH. DNA was treated with exonuclease HI and the reactions terminated at 30 second intervals. DNA fragments of the desired size were treated with the Klenow large fragment, ligated and transformed into DH5aF'. Single stranded DNA was sequenced by the dideoxy chain-termination method (Sanger et al., 1977) with the universal or reverse primers and the remaining sequence was obtained using specific oligonucleotides as primers. The internal Bg/II PCR fragment was sequenced directly in the Bluescript vector in both orientations using a double stranded DNA template with the universal and reverse primers.  Vector constructions  pRaplGal: The 5'  rapl  Hindll  EcoRI 1 kb fragment was isolated from the Bluescript  vector, gel purified and ligated into pDdGall7 (Harwood and Drury, 1990) that had been digested with Xbal, filled in with Klenow fragment and digested with EcoRI. The junctions were confirmed by sequencing.  pVEHRap: The pVEII vector (Blusch et al., 1992) was modified to remove the discoidin  ATG translation start site by digestion with  treatment with E.  coli  Xbal  followed by  DNA polymerase I in the presence of excess dATP in  order to create a single stranded deletion which extended to the first internal adenosine nucleotide. Subsequently the vector was treated with SI nuclease and then the Klenow large fragment of £. coli DNA polymerase I to generate blunt ends before recircularization with DNA ligase. The deletion of the ATG in the modified vector was confirmed by DNA sequencing. This modified vector was then linearized with Kpnl and treated with T4 DNA polymerase to generate a blunt end. An EcoRI fragment containing a full length rapl cDNA was treated with the Klenow large fragment of £.  coli  DNA polymerase I to  fill in the recessed ends, and then ligated into the modified vector in the sense orientation downstream of the  discoidin  promoter, recreating the  flanking EcoRI sites. The junctions were confirmed by EcoRI digestion and by DNA sequencing. pVEII also contains the actin 15-Tn903 resistance cassette (Blusch et al, 1992) as a G418 selectable marker.  Mutagenesis of the rapl cDNA and construction of pVEII vectors: The site-directed mutations in the rapl gene were generated by PCR. Because the D. discoideum  rapl  gene product contains two additional amino-  terminal amino acids and two additional residues located 17 amino acids from the carboxyl terminus compared to the human rapl  and ras gene  products, amino acids have been numbered according to the consensus alignment of Ras proteins to facilitate comparison (Protein sequence alignments were performed with the CLUSTAL module of the PCGENE program). Mutations were introduced by PCR using the 674 bp D.  discoideum  rapl  cDNA EcoRI fragment in the Bluescript vector as a template (Fig. 4) with  the primers shown in Table 1, and either the reverse or universal primer as appropriate. Reaction conditions are described in detail below. Gel purified products of this reaction were subjected to a second PCR in the presence of universal and reverse primer using a  BglR  digested  rapl  cDNA as a template  (this template is disrupted by an internal gap in the rapl cDNA). This PCR selectively regenerated a full length mutated rapl gene. cDNAs mutated at positions 17 and 38 were digested with Hindlll containing the mutation was cloned into  Hindlll  and Nsil and the region Nsil  digested  rapl  in  Bluescript. The cDNAs mutated at positions 61 and 157 were digested with SstI  and Nsil, and the region containing the mutation was cloned into  Nsil  digested  rapl  SstI  in Bluescript. The site-directed mutagenesis was confirmed  by sequencing the entire region which had been amplified by PCR in order to confirm the mutation and check for any additional introduced mutations. The mutated  rapl  genes were then cloned into the pVEII vector  (Blusch et al., 1992) which had been previously modified to remove the discoidin  ATG translation start site as described above. The modified vector  was linearized with Kpnl and treated with T4 DNA polymerase to generate blunt ends. An EcoRI fragment containing the full length rapl cDNA was treated with the large fragment of £. coli DNA polymerase I to fill in the recessed ends and then ligated into the modified vector in the sense orientation downstream of the discoidin  promoter, recreating the flanking  EcoRI sites. The junctions and the site-directed mutations were confirmed by DNA sequence analysis.  First Reaction  Second Reaction  ^ 1 Extension  ^ 2 Amplification  Figure 4. PCR mutagenesis of the rapl gene The first reaction introduces the mutation into the rapl cDNA. The second reaction consists of 2 steps. The first step utilizes the PCR product of the first reaction as a primer with a rapl cDNA containing an internal deletion as a template in an extension reaction which results in a full length cDNA containing the desired mutation.  The second step utilizes two  oligonucleotide primers which are present in molar excess to amplify the full length product. Arrowheads indicate the orientation of the relevant polymerase reaction. Other reactions (not shown) can occur but do not result in as efficient an amplification.  Table 1. Oligonucleotides used for site-directed mutagenesis of the rapl gene  Mutation  a  Oligonucleotide  3  S17N  GTAGGTAAAAATGCATTGACTGTGC  D38E  CCAACCATCGAAGAATCCTACAG  D38N  CCAACCATCGAAAATTCCTACAG  T61Q  G A T A C A G C T G G T C A A G A A C A A 111 A C  F156L  A A G T T A T A C A A A A 111 G T T C  M u t a t e d bases are underlined  Polymerase chain reactions Dried oligonucleotides were dissolved in 30% NH4OH and purified by two cycles of precipitation with n-butanol followed by resuspension in water (Sawadogo and Van Dyke, 1990). The PCR reactions were performed on an Ericomp Twin Block Thermocycler. The PCR universal and reverse primers were 5' CGTTGTAAAACGACGGCCAGT 3' and 5' CAGGAAACAGCTATGAC CATG 3', respectively. The internal 76 bp  BglR BglR rapl  genomic fragment was isolated from  genomic DNA by PCR amplification using the oligonucleotides RapE 5' AAACCAGATGCCTCTTAGAG 3' and JD13 5' AGCTGCAGA(C/A)ATC(G/A/ T)(C/G)(A/C)TTT(G/A)TTAAC 3' (bases in brackets indicate degenerate positions). The reaction consisted of 50 ng of Ax2 DNA, 25 pmoles of each oligonucleotide, 200 uM of each of dATP, dCTP, dGTP and dTTP, 50 mM TrisHC1 pH 8,0.05% Tween 20, 0.05% NP-40 and 0.5 mM MgCl2 and 1 unit of Taq polymerase in a final volume of 50 ul. The amplification protocol consisted of an initial 'hot start' at 94°C for 5 minutes followed by 30 cycles of melting at 95°C for 1 minute, annealing at 47°C for 2 minutes and extension at 72°C for 1.5 minutes, with an additional 5 minute 72°C extension on the final cycle. Site-directed mutagenesis of the rapl cDNA was performed by two consecutive rounds of PCR according to the scheme shown in Figure 4. The oligonucleotides shown in Table 1 were used to introduce the mutations in a reaction with the universal primer (except for mutation F156L where the reverse primer was used) utilizing the rapl cDNA in Bluescript as a template. The first reaction consisted of 200 ng of template, 20 pmoles of each oligonucleotide, 50 uM of each of dATP, dCTP, dGTP and dTTP, 20 mM Tri&: HCl pH 8.4, 50 mM KC1, 0.05% Tween 20, 0.05% NP-40 and 2.0 mM MgCl2, and 1 unit of Taq polymerase in a final volume of 50 ul. The amplification  protocol consisted of an initial 'hot start' at 95°C for 5 minutes followed by 25 cycles of melting at 95°C for 1 minute, annealing at 50°C for 1 minute and extension at 72°C for 1 minutes with an additional 5 minute 72°C extension on the final cycle. The second reaction consisted of 1 ul of the product of the first PCR reaction, 20 pmoles of universal and reverse primers, 20 ng of Bglll digested template, 50 uM of each of dATP, dCTP, dGTP and dTTP, 20 mM Tris-HCl pH 8.4, 50 mM KC1, 0.05% Tween 20, 0.05% NP-40 and 2.0 mM MgCl2, and 1 unit of Taq polymerase in a final volume of 50 ul. The amplification protocol was the same except 30 cycles were performed. Mutations D38N and F156L were generated with Vent polymerase which has greater fidelity than Taq polymerase. The first reaction consisted of 10 ng of template, 20 pmoles of each oligonucleotide, 400 JJM of each of dATP, dCTP, dGTP and dTTP, 10 mM KC1, 20 mM Tris-HCl pH 8.8, 10 mM (NH4)2S04 2 mM MgSC>4, 0.1% Triton X-100 and 1 unit of Vent polymerase in a final volume of 50 ul. The amplification protocol consisted of an initial 'hot start' at 95°C for 5 minutes followed by 20 cycles of melting 95°C for 30 seconds, annealing for 50 seconds and extension at 72°C for 38 seconds. The D38N mutation reaction was annealed at 40°C while the F156L mutation reaction was annealed at 45°C. The second reaction consisted of 2 ul of the product of the first PCR reaction, 20 pmoles of universal and reverse primers, 20 ng of Bg/II-digested template, 400 IIM of each of dATP, dCTP, dGTP and dTTP, 10 mM KC1,20 mM Tris-HCl pH 8.8,10 mM (NH4)2S04,2 mM MgS04, 0.1% Triton X-100 and 1 unit of Vent polymerase in a final volume of 50 ul. The amplification protocol consisted of an initial 'hot start' at 95°C for 5 minutes followed by 20 cycles of melting at 95°C for 30 seconds, annealing at 45°C for 50 seconds and extension at 72°C for 38 seconds.  RESULTS  The  Dictyostelium  discoideum  rapl  gene: isolation of the genomic sequence  and characterization of the promoter region  Introduction The D.  discoideum  rapl  gene is expressed both during vegetative  growth and during development. Steady state rapl mRNA levels increase during aggregation and then following a period of decline during pseudoplasmodial formation, increase again during the formation of the fruiting body (Robbins et al, 1990). During vegetative growth and early development, a rapl single mRNA 1.1 kb in size is detected. After 6-8 hours of development, this mRNA species is replaced by two mRNA species of 1.0 and 1.3 kb (Robbins et al., 1990). The 1.3 kb mRNA is positively regulated in response to pulses of cAMP (Robbins, 1991). To begin to elucidate the complex regulation controlling rapl mRNA expression in D. genomic DNA encoding the  rapl  discoideum,  gene was isolated and sequenced. In  addition, the ability of the upstream untranslated region to promote expression of a reporter gene was tested.  Isolation of rapl genomic DNA Genomic DNA encoding the rapl gene was isolated by constructing and screening a plasmid library. Genomic DNA from D.  discoideum  strain Ax2  was digested with Xbal and Bglll, size fractionated on a sucrose gradient, and the fraction enriched in fragments between 2 and 8 kb in size was cloned into Xbal, BamHI digested Bluescript vector. Southern blot analysis using the  rapl  cDNA as a probe indicated that 2 fragments, 6.6 and 4.0 kb in size, corresponding to the 5' and 3 regions of the rapl gene would be present in this fraction (Fig. 5). Two positive clones which corresponded to the 3' region of the  rapl  gene were identified by hybridization with the rapl cDNA probe.  There were two possible reasons why the 5' region of rapl was not isolated in the first screen. First, Southern blot analysis indicated that the full length probe gave a stronger signal with the 3' region compared to the 5' region (Fig 5). Secondly, given the large size of the fragment to be isolated, it was possible that the fragment was not stable in E.  coli.  To circumvent these potential  problems, the 2-8 kb fraction of genomic DNA was further digested with Hindi and cloned into Hindi, BamHI digested Bluescript vector. One clone containing the 5' genomic region of the rapl gene was identified by hybridization with the EcoRI-Bglll 5' fragment of the rapl-c51 cDNA. The remaining internal 76 bp Bglll fragment was isolated by PCR using two oligonucleotides that corresponded to sequences within the flanking regions as described in the Materials and Methods. The PCR fragment was digested with Bglll releasing a 76 bp fragment which was then cloned into BamHI digested Bluescript vector. The three fragments are designated A, B and C, respectively (Fig. 6).  Nucleotide sequence of the rapl genomic DNA including the 5' region Nested deletions (Maniatis et al., 1989) of fragments A and B were generated with exonuclease III and sequenced as described in the Materials and Methods. Both strands of fragments A and B were sequenced. The short Bglll fragment C was sequenced directly in the Bluescript vector in both orientations. The nucleotide sequence of the rapl genomic clone is shown in Figure 7. The coding region of the gene was found to be divided into two  1  2  3  4  5  F i g u r e 5. Southern blot analysis of D. discoideum rapl g e n o m i c D N A G e n o m i c D N A (6f/g) f r o m A x 2 w a s digested w i t h B g l l l (lane 1); B g l l l a n d H i n d l l l (lane 2); B g l l l a n d X b a l (lane 3); B g l l l a n d H i n d i (lane 4); a n d EcoRI (lane 5). T h e digested D N A was fractionated o n a 0.8% T B E agarose gel and then transferred a n d fixed onto a nitrocellulose m e m b r a n e . p r o b e d w i t h a rapl  c D N A r a d i o l a b e l e d as described i n the materials a n d  methods a n d washed i n 0.1X SSC, 0.1% SDS at 65"C. (kbp) are indicated.  T h e filter w a s  Molecular size standards  Figure 6. Genomic organization of the D. discoideum  rapl  gene.  Exons are shown as hatched boxes. The isolation of fragments (A), (B) and (C) is described in the text. Restriction sites HincII, EcoRI, Bglll and  Xbal  are designated H, E, B and X, respectively. The gap in fragment A indicates a region omitted to compact the figure.  a a c a c t a c a c a a a c a t t g a c acaggcacacacacaaaata a a a a a c c a c c c a c t t a a t t t a a t t t a t t t t t a t t a t t a t t  80  atattttttttttttatttt tttatatttatttttttttt atttatttcaactttttttt tttttttttttttaaaaaaa  160  ataataatattagtgataaa  ccccttcctacaaattaatt  240  tttatttttatttttgtttt tttttttttctttttttttt  320  ttttctttttttttcatttt tattttaattttttttttat ttatttattattactatttt tttttttttatatacatcct  400  tcttaaaaccatttattagt  aagtattttatttgtttttt ttttttttttttttatttat tttattttttttattttttt  480  tttctctatataggatttgt  aatctttttttaatcacttt  aataatatacaattaaaact a t t t t c c c a a a t t t c t t t t t  t t a t a a t a a c c a a t t c a a a a aaaaaaaaaaaaaatttatt  tttatacaagggtaaaaaaa aaaaaaaaaacgatggggag 560  c c a c c t t t t g a t t c a a a a a a ataaaaaaaaaaaaaaaaaa g a a a a a t t a t t t t t t a a a a a  aaaattaaattttttttttt  640  t t t t t t a a t t t t t t t t t t t t ttttccaacKaaaaattttt t t c a t c a c a a a a a t t t t t t t  tttttattttttatttttat  720  acacacatatatatataaca c t t t g t t g t t t t t a t t t a t t tatttaataaccatcaccca accaaattgtaatttaatac  800  attcaactaattaaataatt  880  t t t t t t a a t t a c a t a t t a t a t t t t t t a a a t t a t t a t a a a a ataagaatttaaaaaaaaaa  aaaaaaaaaaaaaaaaaatt a t c t a t t t t c a a a a a a c t t t aaacaagATC CCTCTTOGAG  caaaactcaataaaactaaa t a a a a g g a a a t a t a t a t t a t  960  AATTCMAMX^STCGmTA QGTTCMGTQGTCTAGGTAA ATCTGCrm^TSTOCAAT 1040  TTCTTCAflGGTAXlTlTUl'l' GAAAfiGT^GATCCAACCAT CGMGAITCCTACAGAAAAC AflGTCGAAGTTGAIAGCAftT 1120 CAftTGCftTGTTflGAAfllTrr ASAOaCAGgtaagtttaaat t t t a c t g t t a t a a a g t g a a c a t a a a a t c a t t t t t t t t t t t 1200 aataatgaagaaaaaaaaaa aaaaaaaaatcaaaaaaaaa aaaaaaaaaaaaaaaaaaaa aaaaaaaatatcaaaa-tttt 1280 ttttaaagtggttaattaat  ttaactataactcaatattt  t t t t t t t a a a a a a a t t a t t a gCTOGTflCaXSaft^AarTTflC  attcctttactaatttattt  a t t a t t t t t a t a t a t a t t t t 1360  TQCAATCSflGfiGflTCITrflCA TGAAAAAIQGTCAfiGGTTTT 1440  GJTTTTflCTSffilOTCAMCAT TTOW^TCX^TTTTAflCG ftSTT^VJCAGRICTCCGTGAA CAAATTCTCACIflGTTftflGGA 1520 TTGTGAfiGATCTTCCAA3X3G TTCTTGTTGGTAflCAAAIGC GAaCTCC^aCOiflCGTGT TftriaGCflaV3AftCMGC3TC 1600 iWaAflCTCQCTCGTAAATTT GGTGaTTGTTMTTmaSA AGCATCTQCCAAGAAIAAAG TEARIGTTGAfiCAAATTTTC 1680 TftTAMITAATCCCTCAAAT CAACCGTAAAAAXCAGTTC GTCCftXAfiGCAAflGCTAAA TCAAAATCOXKTTTATTGta 1760 aacaatccatcaactctcca acacccttccatactcaccc  a c c c a t t t c a a a t g t a a c a a ttgaaaaacagaaaaaaaaa 1840  aaaaaaaaaaacaggaaaaa a a a a a a a c a c t t t t t t a a a a aaaaaaaaaaaaataataat a a t a a t a a t a a t a a c c a g t a 1920 atatagtaaatatatatcgt  aaagataccaaaatatgtaa t a a a t a a a t t t t t t t g t t t t t t t t t t a  1987  Figure 7. Nucleotide sequence encoding the rapl gene The initiating ATG is doubly underlined, nucleotides coding for the Rapl protein are in upper case, while flanking and intron sequences are in lower case.  exons separated by a 234 bp intron. The 5' and 3' non-coding sequences and the intron contained a high adenine/thymine content (87%). The nucleotide sequence of approximately 1 kbp of the region upstream of the coding region was also determined.  Analysis of the rapl promoter To determine if the isolated 5' untranslated fragment of the rapl was sufficient to promote expression from a reporter gene, the entire 5' fragment including the first 7 codons of the coding sequence was ligated in frame upstream of the (3-galactosidase reporter gene in the vector pDdGall7 (Harwood and Drury, 1990). Stable G418 resistant transformants were generated using this expression plasmid  (RaplGal)  Materials and Methods. Vegetative Rapl  cells expressed (3-galactosidase  Gal  as described in the  activity whereas the parental Ax2 cells had a very low level of endogenous |3galactosidase activity (Fig. 8). p- galactosidase activity was detected both in vegetative cells and after 4.5 and 9 hours of development and there was a slight increase in p-galactosidase activity at 9 hours. Cells transformed with pDdGall7 had similar levels of p-galactosidase activity to the Ax2 cells (Fig. 8). These results indicate that the 1 kbp upstream region was sufficient to promote expression of p galactosidase.  12  Figure 8. p-galactosidase expression under the control of the 1 kb 5' untranslated region of the rapl gene. Cell free extracts were prepared from the RaplGal the indicated times and from Ax2 cells and  pDdGall  transformant cells at  7 cells. The height of the  bar represents the mean and the error bars represent the standard deviation of P-galactosidase specific activity (nmoles of substrate hydrolyzed/ min/mg of protein) from four determinations for the pDdGall  7  RaplGal  and Ax2 cells. The  cells were analyzed only twice and no error is shown.  Altered morphology of vegetative amebae induced by increased expression of the Dictuostelium  discoideum  rapl  gene  Introduction Rapl proteins have been implicated in diverse roles in various cell types. Kitayama et ah, (1989) isolated a rapl cDNA, Krev-1, based on its ability to suppress the transformed phenotype of K-ras transformed NIH 3T3 cells. Flattened, more adherent cells with reduced tumorigenicity were isolated following fransfection with the rapl cDNA. Rapl also prevents Ras induced germinal vesicle breakdown when microinjected into Xenopus  laevis  oocytes  (Campa et ah, 1991). However, in other cell types, Rapl has effects that are not antagonistic to Ras. Microinjection of RaplB into Swiss 3T3 cells induces DNA synthesis and affects cell morphology (Yoshida et al., 1992). In S. cerevisiae,  the Rapl homologue RSR1/BUD1 is required for control of bud  orientation (Bender and Pringle, 1989), whereas Ras regulates adenylate cyclase activity (Kataoka et al., 1985; Toda et al., 1985). In addition RaplB has been identified in platelets (Siess et al, 1990) and upon platelet activation, it becomes phosphorylated by protein kinase A and associated with the cytoskeleton (Kawata et al, 1989; Fischer et al, 1990). Rapl has also been implicated in the oxidative burst process in B lymphocytes (Maly et al, 1994). In an attempt to identify a role for Rapl in D. discoideum,  the consequences  of expressing high levels of Rapl protein in D. Discoideum were studied.  Effect of Rapl overexpression on cell morphology To determine the effects of high levels of the Rapl protein in vegetative D. repressible  discoideum,  discoidin  the  rapl  promoter. The  cDNA was expressed from a folate rapl  cDNA was cloned downstream of  the discoidin  promoter in the pVEII vector (Blusch et al, 1992) as described in  Materials and Methods (Fig. 9) and the expression vector was introduced into D.  by calcium phosphate precipitate transformation.  discoideum  Transformants were grown either in the presence or absence of folate and increased expression of the Rapl protein was detected by western blot analysis using the anti-Rapl specific antibody. Three of three transformants analyzed showed inducible expression of the Rapl protein (Fig. 10) (The increase in Rapl expression observed in the folate containing samples compared to Ax2 was due to incomplete repression by folate at high cell densities). Preliminary observations indicated that under inducing conditions the Rapl overexpressing transformants had an altered cell morphology not observed in the parental Ax2 cells (See Fig. 25 for AX2 morphology) and also failed to round up after HL5 stimulus (data not shown). Analysis of additional transformants derived independently, showed that 6/7 transformants exhibited an impaired response to HL5 stimulation. A transformant designated  Rapl  was selected for detailed analysis. As a control strain, a  transformant containing pVEII Gal, which expresses (3 galactosidase under the control of the discoidin  promoter, was generated.  To examine the effect of overexpression of the  rapl  gene on cell  morphology, Rapl was plated on a glass coverslip and incubated for 15 hours in conditioned medium or in HL5 with folate.  Conditioned medium  contains a pre-starvation factor (PSF) (Clarke et al, 1987; Clarke et al, 1988), which enhances expression from the discoidin Rapl  promoter (Rathi et  al,  1991).  grown in conditioned medium contained morphologically aberrant  cells which were absent when  Rapl  was grown in HL5 in the presence of  folate (Fig 11). The aberrant cells were flat and spread out with occasional dark regions located around their periphery.  63  Figure 9. The pVEII Rapl expression vector The full length  rapl  cDNA EcoRI fragment was ligated into the  modified pVEII vector in the sense orientation as described in the Materials and Methods, recreating the EcoRI sites (RI). The discoidin  promoter  transcriptional start site is indicated by the arrow. The 5' EcoRI junction sequence and the ATG of the initiating methionine of rapl are underlined.  12 3 4 5 6 7 - 80 -32.5 "*-18.5  Figure 10. Expression of Rapl protein in cells transformed with the pVEII Rapl vector Total protein (10 //g) from Ax2 cells (lane 1) and three independent transformants (lanes 2 and 3; 4 and 5, 6 and 7 respectively ) grown in HL5 medium to a density from 1x10° to 3x10° cells/ml either in the presence of folate (lane 2, 4 and 6) or the absence of folate (lanes 1, 3, 5 and 7) were subjected to SDS-PAGE and transferred to nitrocellulose. The blot was reacted with an anti-Rapl peptide antibody and the signal was detected by ECL as described in the Materials and Methods. The molecular masses of the size markers are indicated in kDa. The level of Rapl protein in Ax2 cells is unaffected by folate or conditioned media (S. Louis and M. Khosla, unpublished observations)  65  cell size  Figure 11. Morphology of vegetative Rapl cells. Dark field micrographs of Rapl cells on a glass coverslip after (A) growth in conditioned medium for 15 hours and (B) growth in the H L 5 medium containing 1 m M folate (B). (C) Comparison of the relative cell areas (n = 54 cells for inducing conditions and n = 81 for non-inducing conditions). Solid bars, cells under inducing conditions; hatched bars, cells under noninducing conditions. Cell size is shown in arbitrary units. Ax2 cells grown in conditioned media have a similar appearance to the Rapl cells grown in the presence of folate (see Fig. 25)  The mean and distribution of the relative cell areas were determined from digitized photographs of uninduced cells and cells induced with conditioned medium. The results from a representative analysis is shown in Figure 11 C. Uninduced cells showed a moderate distribution of cell sizes, while induced cells showed a broader distribution, ranging from normal to highly spread. The difference between the mean areas of cells grown under inducing and non-inducing conditions was statistically significant (p<0.01). Analysis of forward light scatter of Rapl cells growing in suspension under inducing conditions showed changes in the distribution of forward light scatter with a small but significant decrease in the mean (P<0.01) compared to Ax2 cells (Fig 12). These results indicate that although the light scattering properties of the cells are altered, cell volumes are not significantly increased, suggesting that the increased size on plastic is due to a spreading of the cells.  Time course analysis of Rapl protein levels and cell morphology after induction of the discoidin promoter To determine if there was a correlation between levels of Rapl protein and the number of cells with an abnormal appearance, both parameters were analyzed at different times during induction with conditioned medium (Figs. 13 and 14). Western blots probed with an anti-Rapl peptide antibody showed a small increase in the levels of Rapl protein from Rapl cells after 8 hours of induction compared to the folate treated control prior to induction. The levels of the Rapl protein increased further after 16 and 24 hours of treatment with conditioned medium. When the cells were treated with conditioned medium, the number of abnormal cells was significantly increased after 8 hours (p<0.01) and reached a maximum after 24 hours (Fig. 13).  67  A  B  C  D  Figure 12. Forward and side light scatter analysis of vegetative Rapl cells. Ax2 (A and B) and Rapl cells (C and D) were analyzed for forward (xaxis) and side scatter (y-axis) (A and C). Shown is a single parameter histogram plotting cell number (Y axis) versus forward light scatter of Ax2 (B) and Rapl cells (D) in H L 5 .  30% n  Folate  4Hrs  8 hrs  16 hrs  24 hrs  Figure 13. Time course analysis of cell morphology after induction of the discoidin  promoter  Rapl cells were incubated in HL5 medium containing 1 mM folate (folate) or in conditioned medium for the indicated times and the number of cells with a flat spread morphology was determined for 6 separate fields. Between 200 and 600 cells were analyzed for each timepoint. The height of the bar represents the mean and the error bars represent the standard deviation.  F i g u r e 14. T i m e course a n a l y s i s of R a p l protein u p o n i n d u c t i o n of the discoidin  promoter  Total protein (10 //g) from the cells incubated in H L 5 in the presence of folate (lane 1) and in conditioned m e d i u m (lanes 2-5) for 4 h (lane 2), 8 h (lane 3), 16 h (lane 4), and 24 h (lane 5) were subjected to S D S - P A G E and transferred to n i t r o c e l l u l o s e . T h e blot w a s reacted w i t h an a n t i - R a p l peptide a n t i b o d y and the signal was detected by E C L as described in the Materials and Methods. T h e molecular masses of the size markers are indicated in k D a .  Localization of F-actin To determine if the observed morphological changes of the  rapl  transformant cells were associated with alterations of the cytoskeleton, the cells were fixed and stained with FLTC-phalloidin to visualize the distribution of F-actin. Under inducing conditions, abnormal Rapl cells had pronounced peripheral actin staining extending around most of the circumference of the cell (Fig. 15 A to D). The actin staining appeared to coincide either with flattened regions or thin ridges at the periphery of the cell that could be seen in the Nomarski image. In contrast, actin was distributed in clumps at the edges of uninduced Rapl  cells (Fig. 15 E and F). This is consistent w i t h  previous reports that showed that F-actin in vegetative D. discoideum  is  typically punctate in distribution and present in pseudopods during cell translocation (Rubino et al, 1984; Hall et al, 1988).  Effects of Rapl expression on growth The Rapl  ^  cells grew slowly jvvith a doubling time of 12.0 hours when  grown in H L 5 medium without folate compared to a doubling time of 8.2 hours in the presence of folate (Fig. 16). The parental Ax2 cells had a doubling time of 8.6 hours. In addition, it was observed that the Rapl  cells clumped  together when grown in shake suspension at densities between 1 x 1 0 ° and 5x10^ and this cell-cell adhesion could be disrupted by the addition of 10 m M E D T A (data not shown).  A quantitative analysis of the Rapl  cells by the  Gerish laboratory confirmed the observation of increased cell-cell adhesion and demonstrated that the cell-cell adhesion was disrupted by 10 m M E D T A (Fig 17).  71  Of V.  0  >  >  e  IK Figure 15. Localization of F-actin in Rapl cells. Rapl cells on glass coverslips were incubated for 24 hours in conditioned medium (A to D) or in H L 5 medium containing 1 m M folate (E and F). Cells were fixed and stained with FITC-phalloidin and photographed using fluorescence microscopy (A, C and E) and Nomarski optics (B, D and F). Bar = 30/im.  10  8  io -£ 5  0  • 1 • 20  1 • 40  1 60  80  time (hours)  Figure 16. Growth of the Rapl transformant Ax2 (O) or Rapl ( • , • ) cells were washed twice in BS buffer and inoculated into HL5 medium ( O, • ) or P1L5 medium containing 1 mM folate (• ) at a starting density of 5x10^. The line of best fit for cells growing exponentially between 1x10^ and 5x10^ is shown.  0  TO  20,  30  40  50  60 [min]  ' 10  20  30  40  50  60 [min]  B 0.8  0.6 -  0.4 -  0  Figure 17. Cell-cell adhesion properties of the Rapl cells Rapl (V, • )and Ax2 cells ( ° , • ) were (A) grown in HL5 medium to 2xl()6 cells/ ml and (B) developed for 6 hours in shake suspension and then assayed for cell agglutination by measuring light scattering with an agglutometer in the presence ( • , • ) and absence ( V , ° ) of 10 mM EDTA. This data was provided by J. Faix of the Gerish laboratory.  The response of cells to azide treatment It has been shown previously that wild type D.  discoideum  cells round  up and detach when treated with azide, but that the effect of azide is abrogated in cells with a myosin II gene disruption (Patterson and Spudich, 1995). Since the morphological alterations induced by the expression of Rapl are associated with changes in the cytoskeleton (Rebstein et al., 1993), the response of Rapl expressing cells to azide was studied. It was observed that 57 % of wild type Rapl cells remained attached to the substratum after treatment with azide in contrast to the Ax2 cells which almost all detached from the substratum (Fig. 18).  Determination of the number of nuclei in transformed cells The abnormal cell morphology of the activated Rapl cells was similar to that observed for the myosin II heavy chain and profilin mutants that exhibit defects in their ability to undergo cytokinesis (De Lozanne and Spudich, 1987; Knecht and Loomis, 1987; Haugwitz et al, 1994). The number of nuclei in the Rapl transformant cells was therefore determined. There was no increase observed in the number of cells with multiple nuclei. Both the Rapl  transformant and the Ax2 cells had an average of 1.2 nuclei per cell.  Cell motility analysis Since the vegetative  Rapl  transformant cells appeared to have an  abnormal morphology with F-actin distributed around the cell periphery and were also impaired in their ability to regulate their morphology in response to a nutrient signal, the ability of the cells to translocate was evaluated. A field of cells was photographed and then rephotographed after a 30 minute interval (Fig 19). Both normal sized and highly spread Rapl cells were motile  and the distances that they had translocated appeared similar to the distances translocated by the pVEII  Gal  transformant. However, a preliminary, more  quantitative analysis by the Gerish laboratory using computer video analysis, suggested that there may be a slight reduction in the rate of cell translocation (personal communication).  Analysis of morphology after HL5 stimulation When D.  discoideum  cells are transferred from starvation to growth  conditions they rapidly round up and detach from the substratum (Schweiger et al.,  1992; Howard  et al,  1993). To determine whether  Rapl  cells were  capable of this modulation of cell shape, Rapl cells and the parental Ax2 cells were plated on a glass coverslip and starved for 8 hours in BS buffer. The Rapl  cells had a flat and spread appearance after starvation while the Ax2  cells had an elongated appearance characteristic of starved cells (Fig. 20 A and C respectively). The distribution of the relative areas of the starved cells was determined (Fig. 20 E), and the mean of the relative cell areas was significantly larger in Rapl cells compared to the Ax2 parental strain (p<0.01). Cells were then stimulated by replacing the BS buffer with nutrient HL5 medium. After 20 minutes a limited number of Rapl  cells had rounded up,  while all the Ax2 cells had rounded up (Fig. 20 B and D respectively). A time course analysis showed that all the Ax2 cells had rounded up by 10 minutes whereas the response for Rapl cells was strongly inhibited, with 29% of the cells rounded up after 20 minutes (Fig. 20 F).  Figure 18. T h e effect of treating cells w i t h azide A d h e r e n t vegetative A x 2 ( A a n d B), a n d Rapl ( C a n d D ) cells w e r e incubated in conditioned m e d i u m for 24 hours ( A , C,) and then treated w i t h 2 mM  s o d i u m a z i d e i n HL5 for 3 m i n u t e s a n d w a s h e d 3 times to r e m o v e  detached cells (B, D).  Figure 19. The motility of vegetative cells. Rapl  cells (A) and pVEII  Gal  cells (B) were photographed at 30 minute  intervals with dark field optics. The cell outlines were manually traced and the figures superimposed. The red outline indicates the original position of the cell and the blue outline indicates cell position after 30 minutes.  Figure 20 Effect of HL5 stimulation on Rapl and Ax2 cells. Rapl (A, B) and the parental Ax2 cells (C, D) were plated at 3x10^ ' cells / cm2 on glass coverslips, incubated 24 hours in HL5 medium and then starved for 8 hours in BS buffer (A, C). HL5 nutrient medium was introduced for 20 minutes (B, D). (E) The relative cell areas of the cells prior to HL5 stimulation were determined. Solid bars, Rapl cells; hatched bars, Ax2 cells. Cell size is given in arbitrary units (n = 52 cells for Rapl and n = 94 for Ax2, respectively). (F) The time course of cell rounding in response to HL5 stimulation. 40 to 100 cells were counted at the indicated times and the percentage of round Rapl cells ( •) and Ax2 cells ( • ) was determined.  Localization of F-actin in cells treated with HL5 after starvation To observe whether the HL5 stimulus altered the distribution of Factin in cells with a flat spread appearance, the cells were fixed and stained with FLTC-phalloidin. After the reintroduction of HL5 medium, flat spread Rapl  cells still exhibited pronounced peripheral actin staining (Fig. 21 A and  B) which appeared to coincide with a highly flattened region around the edge of the cell that could be seen in the Nomarski image. By contrast the HL5 stimulated Ax2 cells were completely round with localized actin staining (Fig. 21C and D).  Pattern of tyrosine phosphorylation of actin after HL5 stimulation The rounding of starved D . discoideum cells after the reintroduction HL5 medium correlates with a rapid tyrosine phosphorylation of a single major 45 kDa band previously shown to be actin (Schweiger et al, 1992). Since the  Rapl  cells did not respond to the HL5 stimulus, tyrosine  phosphorylation was examined by western blot with an anti-phosphotyrosine antibody (Fig. 22). A dramatic increase in phosphorylation of a single major 45 kDa band (band A) was observed in both Rapl and Ax2 cells within 5 minutes after HL5 stimulation (Fig. 22 A and B, respectively).  This  phosphorylated band increased in intensity for 25 minutes after HL5 stimulation in both Rapl and Ax2 cells. This 45 kDa band corresponded to a prominent band in a Ponceau S stain of total protein on the same nitrocellulose blot and is likely to be actin. These data indicate that although tyrosine phosphorylation of actin may be necessary for cell rounding, it is not sufficient for it to occur. Some minor differences in the pattern of other tyrosine phosphorylated proteins were observed upon starvation of  Rapl  compared to the Ax2 cells (Fig 22 lane S panels A and B, respectively).  Figure 21. Localization of F-actin i n starved and H L 5 stimulated cells. Rapl cells ( A a n d B) a n d the control A x 2 cells (C and D) were fixed 20 minutes after refeeding, stained w i t h F I T C - p h a l l o i d i n a n d then p h o t o g r a p h e d u s i n g fluorescence m i c r o s c o p y ( A a n d C ) a n d N o m a r s k i optics (B a n d D ) . B a r = 30fim.  Figure 22. Protein tyrosine phosphorylation after FIL5 stimulation. Total protein (10 Mg) from Rapl (A) and Ax2 cells (B) were subjected to SDS-PAGE and transferred to nitrocellulose.  The protein was from  vegetative cells (V), cells starved for 8 hours (S) and then exposed to HL5 medium for 5 min (5), 10 min (10), 15 min (15), 20 min (20) and 25 min (25). The blot was reacted with an anti-phosphotyrosine antibody and the signal was detected by ECL as described in the Materials and Methods. The molecular masses of the size markers are indicated in kDa. The letter A indicates the putative actin band.  S 5 10 15 20 25  -32.5 -27.5  These differences have not been analyzed further.  Analysis of the erasure response of Rapl cells Developing wild type cells are not committed to development and the reintroduction of nutrients leads to a return to normal growth. As the Rapl cells did not morphologically respond to the HL5 nutrient medium, it was possible that they would be unable to return to normal growth after initiating development. However, starvation for 8 hours did not affect the viability of the Rapl transformants. Both Rapl and control pVEII Gal transformant cells were fully viable and formed colonies at almost 100% efficiency in a colony formation assay. After nutrients have been reintroduced to cells, they retain the capacity to rapidly recapitulate development if starved again. However, after 2 hours in nutrients, this ability is rapidly erased by a mechanism that involves mRNA degradation (Soil and Waddell, 1975; Waddell and Soli, 1977; Kraft et al., 1989; Chandrasekhar et al, 1990). This 'erasure' response to nutrients was tested in the Rapl strain (Table 2). Rapl and pVEII Gal control transformant cells were allowed to develop in shake suspension for 8 hours and then HL5 medium was reintroduced for either 1 or 2 hours before plating the cells for development on filters. If cells were exposed to HL5 medium for 1 hour, fruiting bodies were observed after an additional 15 hours, while exposure to HL5 medium for 2 hours delayed the onset of the appearance of fruiting bodies by 3 additional hours. The erasure response was identical for both the Rapl  cells and the pVEII Gal control transformant.  Table 2. Erasure of the capacity for rapid developmental recapitulation  3  Rapl  HL5 exposure  pVEII Gal  1  2  1  2  15  +  -  ±  17  +  -  +  19  +  +  +  ±  21  +  +  +  +  Time  a  Transformant cells were starved for 8 hours and then returned to HL5  medium for 1 or 2 hours. The time of the appearance of fruiting bodies after plating cells on filters is indicated. + indicates that part of the population had formed fruiting bodies and + indicates the entire population had formed bruting bodies.  Identification of conserved residues of the Dictyostelium  Rapl  discoideum  protein required to alter cell morphology  Introduction The tumor suppressor activity of human RaplA has been previously analyzed by site-directed mutagenesis and some of the amino acids necessary for the response were identified (Kitayama et al, 1990). Mutations previously shown to activate Ras enhanced tumor suppression by RaplA, and mutations in the effector domain which blocked Ras activation also blocked tumor suppression by RaplA. By contrast, in an analysis of the role of Rapl in oxidative burst in B lymphocytes, both activating and dominant negative mutations had an inhibitory effect, suggesting that in some cases, Rapl is required to cycle between GTP and GDP bound forms (Maly et al, 1994). Increased expression of the Rapl protein in vegetative D. cells from the  discoidin  discoideum  gene promoter correlated with a flattened and spread  cell morphology and an inhibition of morphological responses to external stimuli. To further analyze the role of Rapl in these processes, site-directed mutations of specific amino acids in the D. discoideum  Rapl protein were  generated. Amino acids conserved between the Rapl proteins or conserved between both the Rapl and Ras proteins were selected for mutagenesis and transformants overexpressing these mutated proteins were assessed for changes in their cell morphology and their responses to external stimuli.  The effect of mutated rapl genes on cell morphology Site-directed mutations were constructed in the rapl gene which would encode proteins with following alterations G10V, G12V, S17N, D38E, D38N, T61Q and F156L (the first letter indicating the original amino acid, the  number indicating the position of the amino acid in the protein sequence and the second letter indicating the mutated amino acid). The mutated rapl genes were cloned into the modified pVEII vector as described in the Materials and Methods (Fig. 23) and D. discoideum transformants were generated. None of the transformant clones expressing the G10V or S17N Rapl mutant proteins exhibited an abnormal morphology, while most of transformant clones expressing G12V, D38E, D38N, T61Q, and F156L Rapl mutant proteins contained cells that exhibited the flat and spread morphology (Table 3). To study transformants containing mutated Rapl proteins, a representative transformant which showed a significant proportion of abnormal cells was chosen. For the transformants expressing G10V and S17N mutant proteins (which did not exhibit the abnormal morphology), the transformant clone expressing the highest level of Rapl protein was identified by western blot analysis and selected for further analysis (Fig. 24). The appearance of the selected transformants is shown in Figure 25 and the proportion of cells with a flat spread morphology is shown in Table 4. The transformants F156L  Kapl-GUV,  Rapl-D38N,  Rapl-D38E,  and  Rapl-T61Q  Rapl-  resembled the wild type Rapl transformant (compare Fig. 25, panels B,  D, E, F, and G, respectively with H). In contrast, transformants and Rapl-S17N  Rapl-GlOV  resembled the parental Ax2 cells (compare Fig. 25 panels A  and C respectively with I). The Rapl-G12V,  Rapl-D38E,  Rapl-T61Q  and Rapl-F156L  transformants  (Fig. 26 A, lanes 4, 7, 8 and 9) expressed similar levels of Rapl protein to the wild type  Rapl  transformant (lane 1) while the transformant  Rapl-D38N  expressed somewhat more Rapl protein (lanes 6). As mentioned above,  .86  S17N G12V G10V  D38N D38E  T61Q  F156L  Figure 23. Site-directed mutations of conserved amino acids of Rapl The positions of the introduced mutations are indicated (the first letter indicating the original amino acid, the number indicating the position of the amino acid in the protein sequence and the second letter indicating the mutated amino acid), the hatched boxes indicate regions proposed to be required for guanine nucleotide binding and hydrolysis, and the stippled box indicates the proposed effector domain. The arrow indicates the start of transcription from the discoidin promoter.  Table 3. The effect of site-directed mutations in the rapl gene on cell morphology  Mutation  Clones showing cells with an abnormal cell morphology  3  3  G10V  0/4  G12V  4/4  S17N  0/4  D38E  6/6  D38N  5/5  T61Q  6/7  F156L  9/10  none  4/6  Cells were transformed with mutated rapl genes under the control of the  discoidin  promoter. Independent clones were isolated and examined. The  number of transformants with a flat spread morphology was determined by observing dark field micrographs of 3 independent fields. The transformed clones were categorized as having an abnormal cell morphology if >5% of the cells were flattened and spread.  B  A  1  3  2  4  1  5  - 32.5  H  u  Figure 24.  w  H  w  Expression of  - 27.5 - 18.5  2  3  4 5  - 32.5 - 27.5  I  - 18.5  Rapl protein containing G10V and S17N  substitutions A number of different transformants expressing Rapl proteins with substitutions G10V (A) and S17N (B) were incubated in conditioned medium for 24 hours. Cells were lysed in 1% SDS and 15//g of total cell protein was separated by SDS-PAGE, transferred to nitrocellulose and probed with an antiRapl peptide antibody. Lanes 1-4 represent independent transformants and lane 5 is the parental Ax2 strain. The molecular masses of the size markers are indicated in kDa.  Figure 25. Cell morphology of transformants expressing mutated Rapl protein. Rapl-GWV  D387V (E),  (A), Rapl-GUV  Rapl-T61Q(F),  (B),  Rapl-F156L  Rapl-S17N (G),  (C),  and Rapl  Rapl-D38E  (D), Rapl-  transformants ( H ) and  the parental Ax2 cells (I) were photographed with dark field optics after incubation with conditioned media for 24 hours.  Table 4. The proportion of vegetative cells with a flat spread morphology.  Cell Type Rapl-GlOV  3  3±2  (4)  Rapl-GUV  18 ±4 (2)  Rapl-S17N  2 + 1 (4)  Rapl-D38E  17 + 5 (2)  Rapl-D38N  14 + 5 (2)  Rapl-T61Q  18 ± 3 (3)  Rapl-F156L  17 + 5 (2)  Rapl  18 + 3 (6)  Ax2  a  Flat Spread Cells (%)  3 + 0 (6)  The number of flat spread cells in each transformant was determined by  dark field microscopy of living cells in three separate fields of approximately 50 cells per field. The number of independent experiments is indicated in the brackets. The mean and standard error of the mean are shown.  A  B 1  1 2 3 4 5 6 7 8 9  2 3 4  5 6 7 8 9  -80  -80  -49.5  -49.5  -27.5  -27.5  Figure 26. Rapl protein expression under inducing conditions. (A) Cells were incubated in shake suspension in conditioned medium for 24 hours and then (B) starved for 8 hours. Ax2 (lane 1), Rapl (lane 2), Rapl-GlUV (lane 3), Rapl-G12V (lane 4), Rapl-S17N (lane 5), Rapl-D38N (lane 6), Rapl-D38E (lane 7), Rapl-T61Q (lane 8) and Rnpl-F156L (lane 9) cells were lysed in 1% SDS and 15 //g of total cell protein was separated by SDSP A G E , transferred to nitrocellulose and probed with an anti-Rapl peptide antibody. The molecular masses of the size markers are indicated in kDa  Rapl-GlOV  and Rapl-S17N  were selected based on their expression of high  levels of Rapl protein (lanes 3 and 5).  F-actin distribution in transformed cells The effect of the mutated Rapl proteins on the distribution of F-actin was also evaluated since cells expressing Rapl showed an unusual pattern of F-actin staining.  Rapl-GlOV  and Rapl-S17N  transformant cells were similar  in appearance to Ax2 cells with punctate actin present at the cell periphery while the enlarged F156L  Rapl-GUV,  Rapl-D38E,  Rapl-D38N,  Rapl-T61Q,  Rapl-  and wild type Rapl transformant cells possessed regions of contiguous  F-actin staining at the cell periphery (Fig. 27).  Analysis of morphology after HL5 stimulation As shown in the previous section, Rapl overexpression impaired the ability of cells to change their morphology in response to the reintroduction of nutrients after the onset of starvation. The ability of the mutated Rapl proteins to similarly impair changes in cell morphology was therefore evaluated. Starvation for 8 hours did not appreciably change the levels of Rapl proteins from that observed in cells prior to starvation (compare Fig. 26 panel B with A). The Rapl-GlOV,  Rapl-S17N  and parental Ax2 cells rapidly  rounded up and appeared retractile 5 minutes after the reintroduction of HL5 medium (Fig. 28 panels A, C and I, respectively). In contrast, many GUV,  Rapl-T61Q  and wild type Rapl transformants cells remained flat and  nonrefractile (Fig. 28 panels B, F and H, respectively). The Rapl-D38E, D38N  and  Rapl-  Rapl-F156L  Rapl-  transformants exhibited an intermediate phenotype  with some refractile cells, some non-responding cells and some cells which appeared to contract but not become refractile (Fig. 28 panels D, E and G  E  e  ^|  *  1  1  1=!  U  %  Figure 27. Cell morphology and F-actin distribution. Rapl-GlOV D38N  (E),  (A), Rapl-G12V  Rapl-T61Q  (B), Rapl-SUN  (F), Rapl-F156L  (C), Rapl-D38E  (D),  Rapl-  (G), and Rapl transformants (H) and  the parental Ax2 cells (panel I) were incubated in conditioned media for 24 h, fixed, stained with FITC phalloidin and photographed using fluorescence microscopy as described in the Materials and Methods.  94  9  i  * D  . f  **  Figure 28. The response of starved cells to the reintroduction of HL5 media. Adherent cells on a glass coverslip were treated with conditioned media for 24 hours, starved for 8 hours in BS buffer, and then exposed to HL5 media for 5 minutes. The transformants analyzed were Rapl-GlOV Rapl-GUV  (B), Rapl-SUN  (A),  (C), Rapl-D38E (D), Rapl-D38N (E), Rapl-T61Q (F)  and Rapl-F156L (G), Rapl (H) and the parental Ax2 strain (I).  respectively). To quantitate the response, the proportion of rounded refractile cells was determined 5 and 10 minutes after HL5 stimulation (Fig. 29). The Rapl-T61Q  transformant was not significantly different from wild type  transformant while the  Rapl  transformant exhibited a significant  Rapl-G12V  decrease in the proportion of responding cells (P<0.05). Almost all of the Rapl-GlOV,  and Ax2 cells were round and refractile after 5  Rapl-SI7N  minutes. The Rapl-D38E,  and Rapl-F156L  Rapl-D38N  transformant cells  exhibited an intermediate response, which was less than the Ax2 response but greater than the Rapl transformant response.  The response of cells to azide treatment Wild type D.  discoideum  cells round up and detach when treated with  azide, but the effect of azide is abrogated in cells overexpressing Rapl. GUV  and  Rapl-T61Q  transformant cells were highly resistant to azide  treatment like the wild type Rapl transformant, while the Rapl-GlOV, S17NRapl-D38E  Rapl-  and  Rapl-D38N  Rapl-F156L  Rapl-  transformant cells were  considerably less sensitive to azide treatment (Table 5).  Growth and development The Rapl-G12V,  Rapl-S17N  and the Rapl transformant grew more  slowly than the Ax2 strain in the absence of folate (Table 6). The reduced growth rate did not correlate with increased cell-cell adhesion as the GUV  but not the  Rapl-S17N  Rapl-  transformant exhibited increased cell-cell  adhesion (data not shown). The addition of folate increased the growth rate in all transformants but only the Rapl transformant grew at the same rate as Ax2 under these conditions. All transformants developed normally, both on filters and when plated on bacteria.  > o  LU  >  CO  CM  co  ro Q  Z co CO  Q  CM CO CO  li">  ctf cr  <  Figure 29. The percentage of cells responding to the reintroduction of HL5 medium after 5 and 10 minutes. The number of round refractile cells on three plates was determined 5 min (black bars) and 10 min (hatched bars) after exposure to HL5 medium. The number of cells assessed at each time ranged from 125 to 445. The height of the bar represents the mean and the error bars represent the standard error of the mean from two independent experiments.  Table 5. The percentage of cells remaining adherent after treatment with azide.  Cell type  3  % adherent cells  3  Ax2  3± 1 (3)  Rapl  57 + 10 (3)  Rapl-GlOV  9± 4 (2)  Rapl-GUV  66 + 14 (2)  Rapl-S17N  1± 0 (2)  Rapl-D38E  20 + 11 (2)  Rapl-D38N  12+ 0 (2)  Rapl-T61Q  85 + 15 (2)  Rapl-F156L  24± 6 (2)  Mean + standard error of the mean are shown. The number of experiments is  shown in brackets. Between 200-600 cells were present in the field of view prior to azide treatment.  Table 6. Generation times of transformants expressing mutated Rapl proteins .  Cell type  Ax2  Generation time (hr)  a  + folate  -folate  ND  8.6 (0.98)  Rapl  12.0 (0.92)  8.2 (0.98)  Rapl-GUV  10.4(0.98)  9.6(0.98)  Rapl-Sl 7N  14.3 (0.94)  11.0 (0.98)  Generation times were determined as described in the Materials and  a  Methods for populations between the densities of 1x10^ to 5x10^ cells/ ml since growth within this range was observed to be exponential. The R^ for the line of best fit is shown in brackets.  Activation of the rasG gene alters cell morphology in D.  discoideum  Introduction Microinjection of an activated Ras protein into mammalian cells induces a characteristic transformed cell phenotype (Stacey and Kung, 1984; Feramisco et al, 1984; Bar-Sagi and Feramisco, 1986; Lloyd et al, 1989). The Ras protein induces morphological changes and increases membrane ruffling, reduces stress fiber formation and decreases the formation of focal adhesions (Feramisco et al., 1984; Stacey and Kung, 1984; Bar-Sagi and Feramisco, 1986). These effects of activated Ras on ruffling, stress fibers and focal adhesions are mediated by theras-relatedRac and Rho proteins, which remodel the actin cytoskeleton (Ridley et al, 1992; Ridley and Hall, 1992). D.  discoideum  expresses five  ras  genes during growth and  development (Reymond et al, 1984; Robbins et al, 1989; Daniel et al, 1993b; Daniel et al., 1993a). One of these genes, rasG, is expressed in vegetative cells and encodes a protein which shares 69% overall amino acid identity with the human H-ras gene product (Robbins et al., 1989). Given the effects of activated Ras on mammalian cell morphology and the alteration of cell morphology by the D. discoideum  Ras-related protein Rapl, it was of interest  to determine the effects of "activated RasG on D. discoideum cell morphology.  The effect of RasG protein on D.  discoideum  cell morphology  Transformants generated by M. Khosla (Khosla et al, 1995) with increased levels of RasG, activated RasG-G12T and the putative dominant negative RasG-S17N proteins were analyzed for alterations in the regulation of cell morphology. Under inducing conditions, many  RasG-G12T  cells  exhibited a flattened and spread out morphology with occasional dark regions  around their periphery (Fig. 30 panel B). In contrast, RasG cells exhibited no differences in morphology compared to the parental Ax2 cells (panels A and D). It was observed that 30% of the RasG-G12T cells were abnormally flat and spread out, while 3-4% of the RasG, RasG-SUN and Ax2 cells were somewhat flat and spread out (Table 7). The RasG-S17N cells were more elongated than the Ax2 cells (Fig. 30 panel C). Under inducing conditions, higher RasG protein levels were present in the RasG, RasG-G12T and  RasG-SUN  transformants compared to the parental Ax2 cells (Fig. 31 panel A) with the RasG transformant expressing more protein than the other two transformants. Analysis of forward light scatter of RasG-G12T cells growing in suspension under inducing conditions showed changes in the distribution of forward light scatter with a significant decrease in the mean (P<0.01) compared to Ax2 cells (Fig. 32). These results indicate that although the light scattering properties of the cells are altered, cell volumes are not significantly increased, suggesting that the spread appearance of adherent cells may be due to a change in morphology and not an increase in cell volume.  The localization of F-actin in D. discoideum cells expressing RasG To determine if the observed morphological changes correlated with alterations in the actin cytoskeleton, cells were stained to visualize the distribution of F-actin. RasG-G12T cells with an abnormal morphology exhibited pronounced contiguous peripheral F-actin staining while RasG and RasG-S17N cells exhibited a punctate peripheral F-actin that was characteristic of wild type Ax2 cells (Fig. 33) (see Fig. 27 for Ax2).  F i g u r e 30.  M o r p h o l o g y o f v e g e t a t i v e cells  RasG ( A ) , RasG-GUT  ( B ) , RasG-SUN  (C) a n d A x 2 ( D ) cells  were  p h o t o g r a p h e d w i t h d a r k f i e l d o p t i c s after i n c u b a t i o n w i t h c o n d i t i o n e d m e d i a for 2 4 h o u r s .  Figure 31. Induction of RasG protein expression RasG (lane 1) RasG-GUT (lane 2), RasG-SUN (lane 3) and Ax2 (lane 4) cells were incubated in shake suspension in conditioned medium for 24 hours (A) and then starved in suspension in Bonner's salts buffer for 8 hours (B). Cells were Iysed in 1% SDS and 15 ;/g of total cell protein was separated by S D S - P A G E , transferred to nitrocellulose and probed with an anti-RasG antibody. The molecular masses of the size markers are indicated in kDa.  Figure 32. Forward and side light scatter analysis of RasG-GllT cells. Ax2 (A and B) and RasG-GllT  cells (C and D) were grown in  conditioned media for 24 hours and then analyzed for forward (x-axis) and side scatter (y-axis) (A and C). Shown is a single parameter histogram plotting cell number (Y axis) versus forward light scatter of Ax2 (B) and RasG-GlTT cells (D) in H L 5 . The Ax2 data is reproduced from Figure 12.  Table 7. The proportion of cells with a flat spread morphology.  Cell Type rasG  3  3+1  msG-Gm  30 + 2  rasG-S17N  4±2  Ax2  3  Flat Spread Cells (%)  3+1  The number of flat spread cells was determined in two independent  experiments each consisting of three to five separate fields of approximately 50 cells per field. Cells were viewed with dark field microscopy. The mean and standard error of the mean are shown.  105  B  ^ i j | | i  fj  11  Figure 33. Localization of F-actin in Ras transformed cells Vegetative RiisG cells (A and B), RasG-Gl2T (C and D) and RasG-S17N (E and F) were incubated in conditioned media for 24 hours, fixed, stained with fluorescein isothiocyanate phalloidin and then photographed using either Nomarski optics (A, C, and E) or fluorescence microscopy (B, D and F).  The response of cells to azide treatment Wild type D. discoideum cells round up and detach when treated with azide, but this response is abrogated in cells that overexpress Rapl. Given the fact that the altered appearance and changed F-actin distribution of the G22Tcells resembled that of Rapl cells, the response of the RasG, and  RasG-  RasG-G12T,  transformants to azide treatment was evaluated. As  RasG-S17N  demonstrated previously, Ax2 cells were sensitive to azide treatment and detached from the substratum after azide treatment, while less than 5% of the RasG  and RasG-S17N  34 % of the RasG-GUT  cells remained adherent after this regime. In contrast, cells remained adherent (Fig. 34 and Table 8).  Determination of the number of nuclei in transformed cells The abnormal cell morphology of the activated RasG-Gl2T  cells was  similar to that observed for the myosin II heavy chain and profilin mutants that exhibit defects in their ability to undergo cytokinesis (De Lozanne and Spudich, 1987; Knecht and Loomis, 1987; Haugwitz et al, 1994). The number of nuclei in the  rasG,  rasG-G12T  and  rasG-S17N  transformant cells was  therefore determined. Multinucleate cells were frequently observed in both the RasG and the RasG-G12T  transformants whereas the Ax2 and  RasG-S17N  cells were mononucleate under inducing conditions (Fig. 35). Repression of the discoidin rasG-G12T  promoter by folate abolished the multinucleate phenotype of and RasG transformants (Fig. 35 and Table 9). Both the RasG and  the RasG-G12T  cells had an average of 1.7 nuclei/cell which was significantly  different from the values for Ax2 (P<05).  Figure 34. The effect of treating cells with azide Adherent vegetative RasG (A and B), RasG-GUT  (C and D),  RasG-S17N  (E and F) and Ax2 (G and H) cells were incubated in conditioned medium for 24 hours (A, C , E, and G) and then treated with 2 mM Na azide in HL5 medium for 3 minutes and washed 3 times to remove detached cells (B, D, F and H).  Table 8. The proportion of cells remaining adherent after treatment with azide  Cell type RasG  % adherent cells  3  2 ± 0 (2)  RasG-G12T  34 ± 7 (3)  RasG-S17N  0 + 0 (2)  Ax2  3 + 1 (3)  Approximately 200-600 cells were present in the field of view prior to azide treatment in each trial. The number of independent experiments is indicated in the brackets. The mean and standard deviations are shown.  E  Figure 35. Nuclear staining of Ras transformed cells RasG  (A, D), RasG-GUT  (B), RasG-S17N  (C) and Ax2 (E) cells were  incubated in conditioned media for 24 hours (A, B, C, E) or in HL5 containing ImM folate (D). The cells were fixed and stained with Hoechst dye as described in the Materials and Methods.  Table 9. The number of nuclei per cell  Number of nuclei  3  Strain  Induced  13  Repressed  c  RasG  1.7 + 0.2 (5)  1.1  RasG-GUT  1.7 + 0.1 (5)  1.0 + 0.0 (2)  RasG-SUN  1.1 + 0.0 (4)  1.1  Rapl  1.2 + 0.1 (5)  1.0 + 0.0 (2)  Ax2  1.2 + 0.0 (7)  1.1  (1)  (1)  (1)  Mean and + standard error of the mean of the number of experiments  3  shown in brackets. An average of 200 cells were analyzed per experiment. Cells were grown in conditioned HL5 medium for 24 h.  b  Cells were grown in HL5 medium with 1 mM folate for 24 h.  c  Analysis of morphology after HL5 medium stimulation In view of the similar abnormal morphology of the RasG-G12T and the  Rapl  transformant, the capacity of the cells to respond to the HL5  stimulation was examined. The morphologies of the starved G12T, RasG-S17N RasG-S17N  cells  RasG,  RasG-  and Ax2 cells are shown in Figure 36. The Ax2 cells and the  cells had the elongate morphology characteristic of starved  amebae. In contrast, many of the RasG-G12T  cells retained the extensively  flattened and spread morphology previously observed when induced during vegetative growth. The RasG cells, which had a normal appearance when induced during vegetative growth, became more flattened and spread upon starvation, although not to the same extent as the activated  RasG-G12T  transformant. After starvation for 8 hours, high levels of RasG protein were still present in all three transformants (Fig. 31 panel B). Within 5 minutes of the reintroduction of the HL5 nutrient medium, the Ax2 and RasG-S17N  cells became round and retractile and detached from  the substratum (Fig. 36). In contrast, the majority of the RasG-G12T  cells did  not exhibit this characteristic response although there was some change in their appearance.  RasG  cells did respond to the HL5 stimulus, but not as  rapidly or completely as for the Ax2 strain. The percentage of responding cells was determined 5 and 10 minutes after HL5 stimulation (Fig. 37). It was observed that 59 % of the RasG-Gl  2T  cells failed to respond within 5 minutes  of the reintroduction of HL5 medium and there was only a small additional response after 10 minutes. In contrast, the majority of Ax2 cells and SUN  cells responded within 5 minutes.  response (Fig. 37).  RasG  RasG-  cells exhibited an intermediate  Figure 36. The response of starved cells to HL5 medium. Adherent  RasG  (A and B),  RasG-GUT  (C and D), RasG-SUN  (E and F) and  Ax2 (G and H) cells were incubated in conditioned medium for 24 hours, starved for 8 hours in Bonner's salts (A, C, E and G), and then exposed to HL5 medium for 5 minutes (B, D, F and H).  113  100 i  80 -  _co H> O  o a.  60-  40 -  CO  cr  20  0+"  S17N  Figure 37. The percentage of starved cells that respond to HL5 stimulation. The percentage of round refractile cells was determined 5 min (black bars) and 10 min (hatched bars) after the reintroduction of HL5 medium to starved cells of the indicated transformant strains and Ax2. The height of the bar represents the mean number of round refractile cells and the error bar represents the standard error of the mean for 3 independent plates of cells from a representative experiment. For each determination 200-400 cells were counted.  Expression of Rapl protein in RasG-GUT Rapl  cells and expression of RasG in  cells Given the similar effects of overexpression of activated RasG and Rapl  proteins on cell morphology and the inhibition of the response of cell morphology to external stimuli, western blot analysis was performed to determine if expression of activated RasG resulted in increased Rapl protein levels and likewise if expression of Rapl resulted in increased RasG protein levels. Neither protein caused an increase in expression of the other protein (Fig. 38).  B 1 23 32.527.5-  1 2 3  32.527.518.5  18.5-  Figure 38. Expression of Rapl protein in RasG-G12T cells and expression of RasG in Rapl cells Ax2 (lane 1) RasG-G12T (lane 2), and %>7(lane 3) were incubated in shake suspension in conditioned medium for 24 hours. Cells were lysed in 1% SDS and 15 /ig of total cell protein was separated by SOS-PAGE, transferred to nitrocellulose and probed with a anti-RasG (A) or anti-Rapl peptide antibody (B). The molecular masses of the size markers are indicated in kDa.  GENERAL DISCUSSION Organization and expression of the  rapl gene  DNA sequence analysis of the genomic fragment encoding the  rapl  gene showed that the gene is divided into two exons separated by a 232 bp intron. The position of the intron is not conserved relative to those of the D. discoideum  rasG  and rasD genes (Robbins et  al.,  1992; Reymond et  al.,  1984).  The 5' and 3' non-coding sequences and the intron contain a high adenine/thymine content, which is similar to that for other D.  discoideum  genes (Kimmel and Firtel, 1983). Different  rapl  mRNAs are expressed during vegetative growth and  development (Robbins et al., 1990). During vegetative growth and early development, a single 1.1 kb mRNA is expressed.  After 6-8 hours of  development, this mRNA species is replaced by two mRNA species of 1.0 and 1.3 kb (Robbins et  al,  1990). Since the rapl genomic DNA sequence,did not  reveal any alternative exons, the existence of distinct mRNA species during growth and development must be due either to transcriptional initiation from alternative start sites, to distinct mRNA polyA processing sites, or to a combination of the two. Within the highly adenine/ thymine rich 5' untranslated region of the rapl  gene, there are several clusters of cytosines and guanines that may be  potential transcription regulatory sequences. Cytosine/ guanine rich sequences termed G boxes have been shown to be binding sites for the G box factor (GBF), which is the only D. discoideum transcription factor to be cloned and well characterized to date (Schnitzler et al, 1994). However, GBF mediates activation of cAMP dependent gene expression subsequent to  aggregation (Schnitzler et  al,  1994; Schnitzler et  al,  1995) and since the  rapl  gene is expressed both during growth and development, expression of the rapl  gene is likely to be regulated by additional transcription factors in  addition to possible regulation by GBF. The roles of the cytosine/guanine rich sequences identified in the 5' region of the rapl gene could be determined by analyzing the effects of deletions on expression from the promoter and also by examining the ability of GBF to bind to the promoter. The 5' rapl fragment isolated in this study was tested for the ability to drive expression of p-galactosidase activity in D.  p-galactosidase  discoideum.  activity was detected in vegetative cells and thus the isolated 5' untranslated fragment was sufficient for promoter activity during growth. p-galactosidase activity was also detected after 9 hours of development. As the stability of the Rapl/ p-galactosidase fusion protein in vegetative cells was not studied, it is possible that vegetative p-galactosidase protein may have persisted into development, p-galactosidase activity expressed under the control of the RasD  promoter is detectable in vegetative cells but lost by 4 hours of  development (Esch and Firtel, 1991), suggesting that at least some forms of expressed p-galactosidase enzyme are fairly rapidly degraded during early development, p-galactosidase expressed from the  rasD  promoter is also  moderately unstable during late development (Esch and Firtel, 1991). In contrast, p-galactosidase activity expressed from the cotB,  SP60  and  PsA  promoters is stable when expressed late in development (Detterbeck et al., 1994). The variation in the stability of the p-galactosidase fusion proteins means that the isolated 5' fragment of rapl has not been conclusively shown to promote expression during development.  The effect of Rapl overexpression on D.  High levels of the  D. discoideum  discoideum  cell morphology  Rapl protein were expressed in  vegetative cells in an inducible manner from the discoidin promoter. High levels of the Rapl protein correlated with the appearance of amebae that were abnormally flattened and spread whereas the cells appeared normal when Rapl expression was repressed by folate. It was observed that 18 % of the cells in the population exhibited the abnormal morphology. The cytoskeleton of the flat spread cells was altered with increased F-actin localized to flat lamellipodial regions of the cell periphery. Usually F-actin staining in D. discoideum  is punctate with the most intensely staining regions in the  pseudopods of migrating or chemotaxing cells (Rubino et al., 1984; Hall et al., 1988). Analysis of forward light scatter of cells in suspension suggested that the average cell size was not increased.  Rapl  cells with an abnormal  morphology were still capable of movement, although the rate of cell movement may have been slightly reduced. These results suggest that although the cells were altered in appearance, processes underlying cell motility, such as pseudopod extension and attachment and detachment from the substratum were at most only slightly inhibited. Brief periods of exposure to azide cause D. discoideum cells to contract and detach from the substratum, and high levels of Rapl were found to reduce the response of cells to azide. The mechanism whereby Rapl acts to prevent the response to azide is not known. Azide disrupts respiration, causing a depletion of cellular ATP that is proposed to result in a 'rigor' contraction of the myosin filaments similar to that observed in skeletal muscle (Pasternak et al., 1989). Disruption of myosin II activity in D. discoideum  cells has previously been shown to prevent this response  (Pasternak et al, 1989; Springer et al, 1994; Patterson and Spudich, 1995) and the reduced azide response in cells that overexpress Rapl might be due to a specific alteration of the interaction of myosin II with the actin cytoskeleton. Alternatively, the effect of azide on D. discoideum  cell morphology may  simply reflect the requirements for a continuous supply of ATP in maintaining a normal cytoskeleton (Jungbluth et al., 1994). The effect of Rapl on the azide response may be due to an effect on some other component of the cytoskeleton. The possibility that Rapl acts in some way not related to the cytoskeleton to reduce the sensitivity of the cells to azide also cannot be excluded. HL5 addition to starved D. discoideum  normally causes cells to rapidly  round up and transiently detach from the substratum (Schweiger et ah, 1992; Howard et  al,  1993). In contrast, addition of HL5 to Rapl transformant cells  did not cause them to round up and detach. The inhibition of the response of Rapl  cells to HL5 stimulation was not accompanied by loss of cell viability or  an impaired erasure response, suggesting the cells were neither dying nor unable to revert to vegetative growth. Previously it was shown that D. discoideum cells round up in response to an inhibitor of tyrosine phosphatases, phenylarsine oxide (PAO) (Schweiger et al, 1992), suggesting that the cell rounding response is linked to phosphotyrosine levels.  Furthermore, actin is rapidly tyrosine  phosphorylated in response to HL5 stimulation (Schweiger et al, 1992) and overexpression of tyrosine phosphatase PTP1, but not PTP2, impairs both the cell rounding response and the amplitude and duration of actin phosphorylation (Howard et al, 1993; Howard et al, 1994). However, the  Rapl  transformant showed a normal pattern of actin tyrosine phosphorylation after HL5 stimulation despite its failure to rapidly round up. This result  suggests that 1) the overexpression of Rapl does not block the tyrosine phosphorylation induced by HL5 stimulation; 2) the Rapl cells are still able to detect nutrients and 3) although tyrosine phosphorylation of actin may be necessary (Schweiger et ah, 1992; Howard et al, 1993) it is not sufficient to induce cell rounding in response to HL5 treatment. Given the appearance of the Rapl cells and their failure to respond to azide and HL5, it is possible that the phenotype of flat spread cells was caused by increased cell-substratum adhesion. In mammalian cells, a flatter, more spread cell morphology correlates with increased cell-substratum adhesion (Folkman and Moscona, 1978). However, since the Rapl cells exhibited normal motility, a property which requires the capacity to control adhesion and detachment from the substratum, it is unlikely that increased adhesion was responsible for the altered cell morphology. Rapl  cells exhibited an increase in EDTA-sensitive cell-cell adhesion  when grown in suspension. It is not clear whether this effect was related to the morphological and cytoskeletal changes discussed above. The increase in cell-cell adhesion did not appear to be due to an increase in expression the EDTA-sensitive adhesion molecule gp24 (data not shown) or to increased expression of the EDTA-resistant adhesion molecule gp80 (J. Faix, personal communication.) As detailed in the Introduction, Rapl proteins have been implicated in diverse roles in different cell types and some of these involve cell morphology or cytoskeletal changes. Transfection of a rapl A cDNA has been shown to suppress the transformed phenotype of Ki-ras transformed NTH 3T3 cells, producing flattened, more adherent cells with reduced tumorigenicity (Kitayama et al, 1989), whereas microinjection of RaplB into Swiss 3T3 cells induces membrane ruffling (Yoshida etal, 1992). The yeast Rapl homologue  RSR1/BUD1 is required for the non-random positioning of the bud site (Chant and Herskowitz, 1991), a process which is linked to the spatial organization of polymerized actin. It is interesting to speculate that the effect of high expression of Rapl in D.  discoideum,  which resulted in a flattened  and spread cell morphology, may be similar to the effects observed in the morphological reversion of the phenotype of Ras transformed mammalian cells. Likewise, it is possible that the extended flattened regions observed around the periphery of abnormal D. discoideum Rapl cells may be similar to the lamellipodia that are associated with the formation of ruffles in mammalian cells.  The effects of mutant Rapl proteins on D.  discoideum  cell morphology  Of all the Ras superfamily proteins, the effects of mutations on the protooncogenic Ras proteins have been most extensively studied, so the effects of mutations in amino acids conserved between Rapl and Ras will be discussed initially in the context of the model proposed for Ras (Lowy and Willumsen, 1993; McCormick, 1994; Marshall, 1993). The effects of conserved amino acid substitutions in Ras and Rapl are listed in Figure 10. Briefly, Ras is activated by a switching from a GDP bound state to a GTP bound state and deactivated by hydrolysis of GTP to GDP (Bourne et al, 1990; Lowy and Willumsen, 1993). Consequently, substitutions such as G12V, which result in a constitutively GTP bound protein, cause activation of Ras (Bourne et al, 1990; Lowy and Willumsen, 1993). In addition, substitutions such as F156L, which alter residues not directly involved in nucleotide binding, are also capable of activating Ras, albeit weakly (Quilliam et al, 1995). In contrast, an S17N substitution restricts Ras to a GDP bound  state,  producing  a  dominant  negative  form  of  Ras  Table 10. The effects of conserved amino acid substitutions in Ras and Rapl proteins  Substitution Biochemical Effect  Effect on Ras  Effect on Rapl  G10V  fails to impair activated  hot done  do not bind nucleotides  Ras  binds GTP  activating  1  G12V  2  1  enhances tumor  2  suppression activity S17N  binds GDP  dominant inhibitor  4  dominant inhibitor of  4  oxidative burst  5  D38E  disrupts GAP activity  6  blocks transformation  7  not done  D38N  disrupts GAP activity  6  blocks transformation  7  reduces tumor suppression  T61Q  not applicable  increases rate of GTP  3  reduce tumor suppression  hydrolysis by Rapl  3  Rapl is a substrate for Ras-GAP 8 F156L  structure  1  activating  perturbs protein  disrupts D.  9  9  melanogaster  eye  Clanton et al, 1987; Bourne et al, 1990; Kitayama et al., 1990; Feig and 2  3  4  Cooper, 1988; Maly et al, 1994.; Cales et al, 1988; Stone and Blanchard, 5  6  7  1991; 8 Hart and Marshall, 1990; Quilliam et al, 1995; Hariharan et al, 91. 9  3  10  1  0  (Feig and Cooper, 1988). The S17N mutated Ras protein have been proposed to act by sequestering guanine nucleotide exchange factors, thus blocking the exchange of GDP for GTP on wild type Ras (Feig and Cooper, 1988; Stacey et al, 1991; Farnsworth and Feig, 1991). A G10V substitution prevents the binding of both GTP and GDP (Clanton et al, 1987). Ras activity also requires an effector domain (amino acids 32-40) and substitutions such as D38E or D38N disrupt signal transduction by Ras (Cales et al, 1988; Farnsworth et al, 1991; Stone and Blanchard, 1991) by preventing the binding of effector proteins such as Raf (Vojtek et al., 1993; Warne et al., 1993; Zhang et al., 1993). The Ras superfamily proteins Rho, Rac and Cdc42Hs can be similarly activated or impaired by altering the amino acids equivalent to Ras amino acids G12 and S17, respectively, suggesting that these proteins are also regulated by GDP/ GTP exchange (Paterson et al, 1990; Ridley and Hall, 1992; Diekmann et al, 1991; Ridley et al, 1992; Qiu et al, 1995; Kozma et al, 1995; Nobes and Hall, 1995). Although the effector domains are not highly conserved in the Rho, Rac and Cdc42Hs proteins, mutation of their equivalent domains also blocks activity (Self et al, 1993; Diekmann et al, 1994; Freeman et al, 1994; Zheng et al, 1994). The expression of mutant Rapl proteins, Rapl-GlOV and Rapl-S17N, had no effect on D.  discoideum  cell morphology.  In addition, such  transformants responded to HL5 stimulation and to treatment with azide in the same way as the parental Ax2 cells. These results are consistent with the effects of mutations at position 17 in Ras superfamily proteins which block activity. However, the D. discoideum  S17N Rapl protein did not have an  effect on cell morphology, implying that either the S17N protein did not act in a dominant negative manner, or that interfering with the endogenous Rapl protein activity was not sufficient to alter cell morphology. Although  high levels of S17N Rapl were obtained, it remains possible that further increases in expression would cause an abnormal phenotype. High levels of S17N Ras proteins were required in mammalian cells to cause an effect (Feig and Cooper, 1988; Stacey et al, 1991). The frequency and extent of the abnormal cell morphology was the same in cells expressing the Rapl-G12V protein as in cells overexpressing normal Rapl protein. Similarly, resistance to azide treatment did not differ significantly between the two transformants. However, the cells expressing the Rapl-G12V protein were 14% less responsive to HL5 stimulation than cells overexpressing normal Rapl protein (p<0.05). Together the data suggest that a G12V substitution at best only modestly enhanced the effects of Rapl in D. discoideum.  The same substitution enhances tumor suppression by Rapl  in mammalian cells, but again the effect is not large (Kitayama et al., 1990). This contrasts with the dramatic effect of the same substitution on transformation by Ras (Chang et al, 1982; Stacey and Kung, 1984). Possibly a high proportion of overexpressed Rapl in mammalian and D.  discoideum  cells is in the GTP bound state, and consequently the difference between the wild type and the activated protein is not great. The effects of mutations of amino acids 12 and 17 have been used to infer the requirements for GTP binding on protein activity in Ras superfamily proteins. The tumor suppressor activity of Rapl in Ras transformed mammalian cells was enhanced by a G12V substitution and decreased by a position 17 substitution (although a S17D rather than an S17N substitution was constructed) suggesting that the Rapl tumor suppressor activity required GTP binding (Kitayama et al, 1990). In contrast, both G12V and S17N mutant Rapl proteins, but not wild type Rapl, acted in an inhibitory manner in the oxidative burst process in B lymphocytes, suggesting that in this case it is  necessary for Rapl to cycle between GTP bound and GDP bound states (Maly et al., 1994). This effect on the oxidative burst is reminiscent of the inhibition of Rab-mediated vesicular transport by a non-hydrolyzable GTP analog, which acts to inhibit the shuttling of Rab proteins between two cellular compartments (reviewed in Hall, 1990; Rothman and Orci, 1992). In an analysis of the  S. cerevisiae  Rapl homologue RSR1/BUD1, a G12V activating  mutation was unable to rescue a rsrl/bndl  deletion, again consistent with a  requirement for cycling between a GTP and GDP form of the protein (Ruggieri et al, 1992). The contrasting effects of the Rapl-G12T and Rapl-S17N proteins on D.  discoideum  cell morphology suggest that Rapl in D.  discoideum  acts in  a GTP dependent manner and that Rapl is not required to cycle between a GTP and GDP bound form. Overexpression of Rapl, Rapl-G12V and Rapl-S17N proteins resulted in reduced growth rates compared to the parental Ax2 cells. The reduction in the growth rate could be reversed by the addition of folate to repress the discoidin  promoter. The similar inhibitory effects of all these proteins on  growth contrasted with their markedly different effects on cell morphology. Since wild type, presumptive activating mutant and dominant negative mutant proteins all inhibited cell growth, there is no simple way to reconcile the data with models where Rapl acts in a GTP dependent manner or where Rapl is required to cycle between GTP and GDP bound forms. There was no attenuation of the Rapl-induced cell morphology in cells expressing Rapl-T61Q protein compared to cells overexpressing wild type Rapl protein in any of the assays I used. By contrast, the T61Q substitution reduces the tumor suppressor activity of mammalian RaplA (Kitayama et al, 1990). Residue 61 has been proposed to have at least two functions in mammalian RaplA. It is proposed to be involved in GTP hydrolysis and  contributes to the lower intrinsic rate of GTP hydrolysis in mammalian Rapl relative to Ras (Freeh et al, 1990). In addition, residue 61 contributes to the interaction of RaplA with Ras-GAP. The attenuating effect of the T61Q substitution in mammalian RaplA has been hypothesized to be due an increased rate of GTP hydrolysis, possibly due to the enhanced activity of RasGAP on the mutated RaplA protein (Hart and Marshall, 1990). The lack of any disruptive effects of the T61Q substitution on D. discoideum Rapl activity suggests that increasing the intrinsic rate of GTP hydrolysis is not sufficient to reduce the effect of Rapl. In addition, the different effect of the T61Q substitution on D.  discoideum  Rapl compared to mammalian Rapl suggests  that substitution of position 61 may not impair  D.  discoideum  Rapl  interaction with the presumptive GAPs that have yet to be characterized. In this regard, bovine Rapl-GAP (GAP3) activity is not inhibited by a T61Q RaplA substitution (Maruta et al, 1991). Expression of a Rapl protein containing the F156L substitution in D. discoideum  resulted in cells with an abnormal cell morphology. However,  the Rapl-F156L  cells were more sensitive to treatment with azide compared  to Rapl cells and also exhibited a reduced inhibition of the response to HL5 stimulation compared to wild type  Rapl  cells. These data suggest that the  Rapl protein with a position F156L substitution has reduced activity compared to the wild type Rapl protein. This result contrasts with the weakly activating effect of this substitution in Ras and the dominant gain-of-function of this substitution in D.  melanogaster  Rapl (Hariharan et  al,  1991; Quilliam  et al, 1995). The basis of this difference is not known. Although position 38 mutations disrupt transformation by Ras and tumor suppression by Rapl (Sigal et al, 1986; Stone and Blanchard, 1991; Kitayama  et al.,  1990), Rapl-D38E  and Rapl-D38N  transformants cells  exhibited the same abnormal cell morphology as the wild type transformant. However, the Rapl-D38E  and Rapl-D38N  Rapl  cells were sensitive  to azide treatment and they had a phenotype intermediate between that of the parental Ax2 strain and the Rapl-D38N Rapl-D38E  Rapl  transformant after HL5 stimulation. The  cells had a smaller inhibitory effect on HL5 stimulation than the cells, despite the fact that the  Rapl-D38N  cells expressed  somewhat more Rapl protein. This result is consistent with the fact that a D38N substitution is a more substantial change, substituting a basic side chain for an acidic side chain, than is D38E, which only introduces a single additional methyl group. Since these substitutions were in the core region of the effector domain that is conserved in all Ras and Rapl proteins (Bourne et al, 1991), it was anticipated that these mutated Rapl proteins would be inactive.  The partial activity of the Rapl proteins with position 38  substitutions suggests that effector domain requirements for Rapl-induced changes in  D. discoideum  cell morphology differ from those required for  tumor suppression by mammalian Rapl and transformation by Ras. Mutations D38E, D38N and F156L all attenuated the effect of Rapl expression on the response of cells to azide and the response of starved cells to HL5 stimulation, yet did not diminish the effect of Rapl on cell morphology. This raises the question of whether the effect on morphology, the response to azide and the response to HL5 stimulation represent the same or different phenomena. The response of Ax2 cells to HL5 and azide both involve a process of rapid cell contraction and detachment from the substratum in response to a stimulus, although it is not clear that the two responses involve identical cytoskeletal changes. Analysis of cells expressing mutant Rapl proteins showed a good correlation between resistance to azide treatment and the inhibition of the response to HL5 stimulation (Fig. 39), which suggests that these two assays  100% -  80%  60%  40%  20%  0%  Figure 39. Comparison of azide treatment with the HL5 stimulation assay The data for the proportions of cells resistant to azide treatment (open bars) is from Tables 5 and 8 and the data for HL5 stimulation for 10 minutes (black bars) is from Figures 29 and 37. Transformants have been arranged according to the extent of the inhibition of the response to HL5 stimulation. The height of the bar indicates the mean and the error bar indicates the standard error of the mean.  may measure the same effect. The relationship between the flat spread morphology on the one hand and the azide and HL5 stimulation response on the other is more complex. An altered cell appearance and an inhibition of the responses to azide and HL5 stimulation were observed in cells expressing either activated RasG-G12T, Rapl, Rapl-G12V or Rap-T61Q proteins. However, the presence of flat, spread cells in transformants expressing similar levels of mutated Rapl-D38E, Rapl-D38N or Rapl-F156L protein did not predict the response to HL5 stimulation or the response to azide, suggesting that the flat spread cell phenotype is distinct from the two other responses. Analysis of additional mutations in the Rapl effector domain might clarify the relationship between the flat spread cell morphology and the responses to azide and HL5 stimulation.  The role of R a s G in D. discoideum  Under inducing conditions, many of the RasG-G12T cells exhibited an abnormal morphology: they appeared flattened and spread with increased Factin located around the cell periphery, although there was no increase in cell volume as determined by an analysis of forward light scatter (this thesis) and Coulter counter analysis (G. Weeks, personal communication). By contrast, RasG  cells had an appearance similar to that of Ax2 cells. Activated Ras has  previously been shown to have a dramatic effect on mammalian cell morphology (Stacey and Kung, 1984; Feramisco et al, 1984; Bar-Sagi and Feramisco, 1986; Lloyd et al, 1989). Mammalian cells exhibit a characteristic transformed phenotype; becoming refractile, exhibiting increased membrane ruffling and a loss of stress fibers. Ras proteins have also been shown to regulate cell morphology in yeasts. In Schizosaccharomyces  pombe,  Rasl is  required for normal cell shape, in a process involving cdc42sp a Rho-like protein (Chang et  al,  1994) whereas in  Saccharomyces  cerevisiae  RAS2 is  involved in pseudohyphal growth, a process characterized by unipolar budding and an altered cell morphology (Gimeno et al, 1992). Expression of activated RasG-G12T increased the number of D. discoideum  cells that remained adherent to the substratum after azide  treatment and substantially inhibited the cell rounding response to HL5. By contrast, high expression of wild type RasG protein modestly inhibited the response to HL5 stimulation but had no effect on vegetative cell morphology or on the azide response. These results are also consistent with the G12V alteration causing an activation of RasG. The abnormal appearance of the RasG  cells following starvation and the inhibited response to HL5 suggested  that starved cells may be more sensitive than vegetative cells to the effects of overexpression of wild type RasG. However starved RasG cells, unlike  RasG-  G12T cells, could develop normally (M. Khosla, personal communication). It is also worth noting that the effect of RasG overexpression on the azide and HL5 responses contrasted with the results of the Rapl mutant proteins, and was the only indication that the two responses might not be affected equally. Cells expressing RasG-S17N protein were more elongated than Ax2 cells whereas the  RasG  cells appeared normal. RasG-S17N may act as a  dominant negative form of the protein causing an inhibition of the normal activity of the RasG protein as described previously for mammalian Ras S17N protein (Feig and Cooper, 1988). The contrasting effects of the G12T and S17N substitutions suggest that RasG affects vegetative cell morphology in a GTP dependent manner. No increased rate of rounding in response to HL5 stimulation could be detected in RasG-S17N  cells (data not shown). This may  be due to the difficulty of observing a response that is greater than the already  large response of the parental Ax2 cells, or alternatively, as discussed for Rapl, the effect on HL5 stimulation and azide may be separate from the effect on cell morphology. Multinucleate cells were observed in both the RasG and the  RasG-G12T  transformants. However, expression of activated RasG did not increase the proportion of multinucleate cells or the average number of nuclei per cell compared to overexpression of wild type RasG. The similar effects of the two proteins contrasts with the enhanced effect that expression of the RasG-G12T protein had on cell morphology, suggesting that the effect on cytokinesis may be unrelated to changes in cell morphology. Furthermore, overexpression of Rapl altered cell morphology but had no affect on cytokinesis, again suggesting that the two phenomena are not related. Cells with a disruption of the myosin II heavy chain or the profilin genes are also multinucleate and have a flat spread cell morphology (Knecht and Loomis, 1987; De Lozanne and Spudich, 1987; Haugwitz et ai, 1994). However, the consequences of overexpressing RasG or an activated RasGG12T protein differed from those reported for cells disrupted in myosin II or profilin in two respects. First, the proportion of multinucleate cells was not as great and number of nuclei per cell was considerably lower for the and the RasG-G12T  RasG  transformants. Secondly, disruption of the myosin II or  profilin genes also results in cells which are unable to grow in suspension (Knecht and Loomis, 1987; De Lozanne and Spudich, 1987; de Hostos 1993; Haugwitz et al, 1994). By contrast, RasG and RasG-G12T  etal,  cells do grow in  suspension, albeit at a reduced rate (M. Khosla, personal communication).  H o w does overexpression of R a p l a n d RasG-G12T affect D. discoideum cells?  In contrast to the apparently antagonistic roles that Rapl and Ras proteins play in mammalian cells, there are a number of striking similarities between the effects of Rapl and activated RasG overexpression on D. discoideum cells. Expression of either Rapl or activated RasG-G12T caused  cells to become flattened and spread with a redistribution of F-actin around the cell periphery. In addition, expression of both Rapl and activated RasG reduce cell detachment after treatment with azide, and inhibit the rounding up of starved cells in response to a nutrient stimulus. However there are also significant differences between the actions of Rapl and RasG proteins. One difference is that overexpression of wild type and activated Rapl have similar effects on cell morphology and the responses to azide and HL5, whereas all three properties are more affected by expression of activated RasG than by overexpression of wild type RasG. Another difference is that RasG affects some processes which are unaffected by Rapl. Activated RasG expression blocks aggregation (M. Khosla personal communication) whereas wild type and activated Rapl have no effect on development.  In addition,  overexpression of either RasG or RasG-G12T but not Rapl protein resulted in a defect in cytokinesis. These results raise two questions for discussion. First, how does the overexpression of Rapl and RasG-G12T affect cell morphology, and the responses to azide and nutrient stimulation; and second, do Rapl and RasG act on the same or different regulatory pathway (s)? By analogy with the effects of Ras on morphology in mammalian cells, there are two non-mutually exclusive ways for Rapl and RasG to affect D discoideum cell morphology. Overexpression of Rapl and activated RasG-  G 1 2 T may act to disrupt a signal transduction pathway controlled by Rapl and  RasG that directly regulates the cytoskeleton. In mammalian cells, Ras affects actin-based processes via pathways involving Rho and Rac, as described in the Introduction. Several Rac and Rho genes have been isolated from D. discoideum  (Bush et  al,  1993a; Bush et  al,  1993b), and it is possible that there  is a network of Ras superfamily proteins regulating D.  discoideum  cell  morphology. One cytoskeletal process that might be regulated by such a pathway is the contraction of the cytoskeleton by myosin II, since this process responds to azide. The assembly of myosin II onto the cytoskeleton is induced by stimuli such as cAMP and is regulated by phosphorylation (Berlot et al, 1985; Berlot et al, 1987; Egelhoff et al, 1993), and overexpression of Rapl and RasG-G12T could disrupt such a process. However, there are probably other signal transduction pathways that directly regulate the cytoskeleton that could be regulated by Rapl and RasG. The second way that overexpression of Rapl and RasG might act is to disrupt the transduction of a signal controlling gene expression which consequently results in an altered cell morphology. Activated Ras transduces a signal to the nucleus which regulates mammalian gene expression (reviewed in Gutman and Wasylyk, 1991; Lowy and Willumsen, 1993) and reduced expression of numerous actin associated genes has been observed in transformed mammalian cells, providing a precedent for such a proposal (Button et al, 1995; Janmey and Chaponnier, 1995). Recently, it has also been shown that other Ras superfamily proteins, Rho, Rac and Cdc42Hs, transduce signals that activate transcription (Minden et al, 1995; Coso et al, 1995); Hill, 1995 #497. There is currently is no direct evidence for activation of gene expression by a Rapl-specific pathway, but the Observation that microinjection of Rapl can stimulate mitogenesis in Swiss 3T3 cells (Yoshida et al, 1992) is consistent with a role for Rapl in transducing a signal to the  nucleus. Since the effects of the Rapl and RasG proteins in this study were analyzed 24 hours after the induction of the discoidin  promoter, sufficient  time had elapsed to allow additional gene expression. discoideum  Candidate D.  genes that might be regulated by Rapl and RasG include those  identified by gene disruption studies that cause similar alterations in cell morphology, such as profilin or coronin (Haugwitz et al., 1994; de Hostos et al., 1993). Actin and myosin II heavy chain levels did not appear to be altered in the Rapl and  RasG-G12T  cells (data not shown).  Do Rapl and RasG act on the same of different regulatory pathway(s)? Given the very similar effects of overexpression of Rapl and activated RasG proteins, it is possible that the two proteins act on the same pathway. The additional effects of activated RasG-G12T on cytokinesis and aggregation could be due to another RasG-specific pathway, or to RasG acting upstream of Rapl and transducing two or more signals, only one of which is subsequently mediated by Rapl. The latter arrangement would be similar to the branched organization of Ras superfamily proteins involved in the regulation of mammalian cell morphology. Alternatively, Rapl and RasG could regulate two different pathways that converge on the cytoskeleton, perhaps at different regulatory sites, but with a similar phenotypic outcome. The relationship, if any, between Rapl and RasG might be addressed by coexpression experiments utilizing dominant negative forms of the proteins; for example, if one assumes that Rapl acts downstream of RasG then expression of Rapl-S17N should block some of the effects of activated RasG-G12T. The above discussion assumes that overexpression of Rapl and RasGG12T disrupts a signaling pathway regulated by the endogenous Rapl and RasG proteins. Although expression of Rapl-S17N did not affect cell morphology, a role for endogenous RasG protein in directly regulating cell  morphology is supported by the observation that the both RasG-G12T and the dominant negative RasG-S17N protein affected cell morphology. However, it is possible that Rapl and RasG-G12T disrupt a pathway that is normally regulated by another member of the Ras superfamily. It is conceivable that Rapl and activated RasG-G12T affect a pathway regulated by RasB or RasC, which are both expressed at significant levels during growth in D. discoideum (Robbins et al., 1989; Daniel et al., 1993a; Daniel et al., 1993b). Rapl and activated RasG-G12T are less likely to affect pathways regulated by other Ras superfamily proteins, such as Rac or Rho that share less amino acid identity with Ras. It has been proposed, based on studies in mammalian cells, that overexpression of Rapl may act in a manner distinct from that of an activated Ras protein. Rapl was proposed to revert the transformed phenotype of Ras transformed mammalian cells by competing for Ras effectors (Freeh et al, 1990; Zhang et  ah,  1993) and it is possible that D .  discoideum  Rapl could  similarly sequester effector or regulatory factors of other Ras superfamily proteins and switch on or off the pathways regulated by these molecules. Since Rapl does not affect cytokinesis and development, whereas RasG-G12T does affect these processes, it is unlikely that Rapl is activating RasG by sequestering negative regulatory factors. However, the effects of RasB and RasC on cell morphology are not known and it is possible that Rapl may act by stimulating or disrupting the effects of these or other as yet uncharacterized Ras proteins. 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