Activated Camp Receptors Switch Encystation Into Sporulation

Activated cAMP receptors switch encystation into sporulation

Yoshinori Kawabea, Takahiro Moriob, John L. Jamesa, Alan R. Prescottb, Yoshimasa Tanakab, and Pauline Schaapa,1

aCollege of Life Sciences, University of Dundee, Dundee, Angus, DD15EH, United Kingdom; and bGraduate School of Life and Environmental Sciences, University of Tsukuba, Ibaraki, 305-8572, Japan

Edited by Peter N. Devreotes, Johns Hopkins University School of Medicine, Baltimore, MD, and approved March 12, 2009 (received for review February 13, 2009) Metazoan embryogenesis is controlled by a limited number of in the most derived group 4 (4). During D. discoideum devel- signaling modules that are used repetitively at successive development, the deeply conserved intracellular messenger cAMP opmental stages. The development of social amoebas shows sim- has multiple roles as a secreted signal, detected by 4 homologous ilar reiterated use of cAMP-mediated signaling. In the model cAMP receptors (cAR1–4) (5). cAMP pulses coordinate the Dictyostelium discoideum, secreted cAMP acting on 4 cAMP recep- aggregation of starving cells and organize the construction of tors (cARs1-4) coordinates cell movement during aggregation and fruiting bodies with a highly regulated pattern of spores and stalk fruiting body formation, and induces the expression of aggrega- cells. Secreted cAMP also up-regulates expression of aggregation and sporulation genes at consecutive developmental stages. tion genes, induces expression of spore genes, and inhibits stalk To identify hierarchy in the multiple roles of cAMP, we investigated gene expression (6). cAR heterogeneity and function across the social amoeba phylog- Single cAR genes were previously detected in 3 more basal eny. The gene duplications that yielded cARs 2-4 occurred late in dictyostelid taxa, but were only expressed after aggregation. The evolution. Many species have only a cAR1 ortholog that duplicated nonhydrolyzable cAMP analog SpcAMPS, which inhibits cAR- independently in the Polysphondylids and Acytostelids. Disruption mediated pulsatile cAMP signaling, disorganized fruiting body of both cAR genes of Polysphondylium pallidum (Ppal) did not formation in these taxa, suggesting an ancestral role for pulsatile affect aggregation, but caused complete collapse of fruiting body

cAMP signaling in fruiting body morphogenesis (7). However, it EVOLUTION morphogenesis. The stunted structures contained disorganized remains unresolved whether cARs also mediated gene expres- stalk cells, which supported a mass of cysts instead of spores; cAMP sion in basal taxa, for which SpcAMPS is a normal agonist, and triggered spore gene expression in Ppal, but not in the cAR null whether the detected cARs were unique or members of larger mutant, explaining its sporulation defect. Encystation is the sur- gene families. vival strategy of solitary amoebas, and lower taxa, like Ppal, can To resolve these questions, we first mapped cAR heterogeneity still encyst as single cells. Recent findings showed that intracellular in the Dictyostelia, showing multiple independent events of cAR cAMP accumulation suffices to trigger encystation, whereas it is a gene duplication. Second, by successively disrupting all cAR genes complementary requirement for sporulation. Combined, the data of the early diverged taxon Polysphondylium pallidum (Ppal), we suggest that cAMP signaling in social amoebas evolved from greatly expanded the opportunities for cAR functional analysis. cAMP-mediated encystation in solitary amoebas; cAMP secretion in Loss of 1 cAR caused reduced fruiting body branching, whereas loss aggregates prompted the starving cells to form spores and not of both disrupted fruiting body formation. Strikingly, without cysts, and additionally organized fruiting body morphogenesis. cARs, Ppal cells could not express spore genes, and formed cysts cAMP-mediated aggregation was the most recent innovation. instead of spores in the stunted structures. Encystation is the major stress response of solitary protists, which is retained in several early developmental signaling ͉ evolution of multicellularity ͉ Dictyostelia ͉ diverging social amoebas. Our data show that cysts are ancestral to Amoebozoa spores, and that activated cARs determine the choice between the 2 developmental pathways. he evolution of novel morphological features is due to Results Tchanges in the developmental signaling processes that shape these features. The number of different signals that shape cAR Heterogeneity in the Dictyostelia. D. discoideum has 4 homol- complex embryos, such as mammals, is surprisingly limited ogous cARs with different functions in chemotaxis and gene because the same signals, such as members of the wingless/wnt, regulation (5). To understand how these functions evolved, we hedgehog, TGF-␤, and FGF families, are used many times over first mapped patterns of cAR gene duplication across the Dic- at successive developmental stages (1–3). Also, the signals and tyostelid phylogeny. Two to 5 test species were selected from their associated transduction pathways are deeply conserved in each of the 4 groups of Dictyostelia (Fig. 1A). Amplification of evolution, often having homologous roles in shaping the body plan of lower invertebrates. The first multicellular organisms Author contributions: Y.K., Y.T., and P.S. designed research; Y.K. and J.L.J. performed most likely deployed preexisting signaling systems from their research; T.M. and A.R.P. contributed new reagents/analytic tools; Y.K. and P.S. analyzed unicellular ancestors, which were used to find food or mates, or data; and Y.K. and P.S. wrote the paper. to evade stress. To fully understand developmental signaling, it The authors declare no conflict of interest. is of fundamental importance to identify which protist signaling This article is a PNAS Direct Submission. pathways were used, and how they were adapted and elaborated Data deposition: The sequences reported in this paper have been deposited in the GenBank to generate the ever increasing complexity of multicellular database [accession nos. EU797651 (DroscAR3), EU797652 (DroscAR4), EU797653 (Dmuc- organisms. cAR1), EU797654 (DmuccAR2), EU797655 (DmuccAR3), EU797656 (DmuccAR4), EU797657 The dictyostelid social amoebas offer unique opportunities to (PviocAR), EU797658 (DrhicAR), EU797668 (PpalTasB), EU797661 (PpseTasA), EU797662 (PpseTasB), EU797663 (DgloTasA), EU797664 (DgloTasB), EU797660 (AanacAR), EU797665 address this issue. They are as genetically diverse as animals, but (AsubcARE), EU797666 (AsubcARF), EU797667 (AsubcARG), EU797659 (DbifcAR), and alternate a sophisticated program of multicellular morphogen- EU797669 (DaurcAR)]. esis with a free-living amoeboid lifestyle. A robust molecular 1To whom correspondence should be addressed. E-mail: [email protected]. phylogeny is available, showing subdivision of all known species This article contains supporting information online at www.pnas.org/cgi/content/full/ into 4 groups, with the model Dictyostelium discoideum residing 0901617106/DCSupplemental.

www.pnas.org͞cgi͞doi͞10.1073͞pnas.0901617106 PNAS ͉ April 28, 2009 ͉ vol. 106 ͉ no. 17 ͉ 7089–7094 Downloaded by guest on September 25, 2021 Fig. 1. Identification and phylogeny of cAMP receptors. (A) Species selection. The published SSU rRNA based phylogeny of all known Dictyostelia (4) is schematically represented. The position of species from each of the 4 major taxon groups that were selected for cAR identification are indicated. (B) Identification and phylogeny of cARs. Partial 324–330-bp cAR sequences were amplified by PCR from genomic DNAs of the test species. Complete coding seqeunces for PpalTasB and DaurcAR were obtained by inverse PCR with primers complementary to the first PCR product. All derived partial and complete cAR amino acid sequences were aligned with previously published cAR sequences, the Ddis cAR-like receptor CrlA and with cAR homologs in nondictyostelid organisms. Phylogenetic relationships between sequences were determined by Bayesian inference (36), from 108 (PCR products) to 253 (full-length protein) well-aligned positions. The Bayesian inference posterior probabilities of nodes are indicated by line thickness. Numerical values are displayed in Fig. S2. The tree was rooted on DdisCrlA and the 3 nondictyostelid receptors. GenBank accession numbers for published sequences are: Arabidopsis thaliana (Atha)GPCR: NP࿝175261; Physcomitrella patens (Ppat)GPCR: XP࿝001765654; Danio rerio (Drer)GPCR: XP࿝001332705; DdiscAR1: P13773; DdiscAR2: AAB25436; DdiscAR3: AAB25437; DdiscAR4: Q9TX43; DmincAR: AAS59250, PpaltasA: BAA99285; DfascAR: AAS59252.

cAR genes from genomic DNAs of the selected species yielded most similar Ppal TasA or TasB. The 3 Asub cARs are quite 1 or multiple products ranging from 324 to 330 bp. Their derived diverse, but still more similar to cARs than to the Ddis cAR-like amino acid sequences shared 59–97% sequence identity between protein CrlA. All nongroup 4 cARs are more similar to Ddis- each other and DdiscARs (Fig. S1A). Four cAR homologs each, cAR1 than to DdiscARs2-4. These data indicate that the gene were amplified from the group 4 species Dictyostelium mu- duplications that gave rise to cARs2-4 occurred in Group 4 taxa, coroides (Dmuc) and Dictyostelium rosarium (Dros). Ppal and 2 and that independent cAR gene duplications occurred at least 3 related Group 2 species, excluding the Acytostelids, yielded 2 more times. cAR homologs each. Acytostelium subglobosum (Asub) yielded 3 cAR homologs, but only a single cAR was isolated from Acy- Expression Patterns of cARs in Ppal. We planned to use the large tostelium anastomosans (Aana). All tested Group 1 and 3 species, genomic fragments of the Dmin and DaurcARs and PpalTasB to and the group-intermediate species Polysphondylium violaceum generate constructs for expression analysis and gene disruption. (Pvio) contained only a single cAR homolog. However, we have not been able to achieve stable transformation We next isolated full-length cAR genes by inverse PCR from of Dmin and Daur cells yet. Ppal can be transformed; therefore, at least 1 species in each group. Full-length sequences were we concentrated on analyzing cAR function in this species. We already available for DdiscARs 1-4 in group 4 (5), Dictyostelium first compared the temporal expression patterns of TasB and minutum (Dmin) cAR in group 3 (7), and 1 Ppal cAR, named TasA. Fig. 2A shows that TasB expression is first visible after 2 h TasA in group 2 (8). Complete coding regions were amplified for of starvation, to reach a plateau at 8 h when aggregation is the Dictyostelium aureo-stipes (Daur) cAR in group 1, and for the completed. TasA mRNA first appears after aggregates have second cAR of Ppal, which was named TasB. Alignment of the formed. Both genes remain highly expressed until late fruiting deduced amino acid sequences of DaurcAR and PpalTasB with body formation. To visualize the spatial pattern of TasB expres- other complete cARs shows that the regions that contain the 7 sion, its 2.3-kb promoter was fused to the LacZ reporter gene and transmembrane helices are highly conserved, whereas the car- transformed into Ppal cells. Developing structures were incu- boxyl-termini of the cARs are very diverse (Fig. S1B). bated with X-Gal to visualize ␤-galactosidase activity. TasB was An alignment of all PCR products and full-length cARs was first expressed weakly at the center of streaming aggregates (Fig. used to assess phylogenetic relationships between the cARs. Fig. 2B). Expression then increased throughout the newly formed 1B shows that the 4 cARs of Dmuc and Dros are most similar to sorogen, but soon appeared strongest at the tip region (Fig. 2 C cARs1-4 in Ddis. The 2 cARs that were identified in Dictyoste- and D). Ppal fruiting bodies form whorls of side branches from lium gloeosporum (Dglo) and Polysphondylium pseudocandidum cell masses that pinch off from the rear of the sorogen. TasB (Ppse), 2 species that are closely related to Ppal (Fig. 1A), are showed high expression at the site of separation of these cell

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B CD F G

E F H I Fig. 3. Phenotypes of single and double tasAϪ and tasBϪ mutants. The Ppal tasAϪ mutant was created earlier (8), and the generation of tasBϪ, tasAϪtasBϪ,

and tasAϪtasBϪ/TasB mutants is outlined in Materials and Methods. Wild-type EVOLUTION (WT) and mutant cells, harvested from bacterial growth plates, were plated on water or charcoal agar, and incubated at 22 °C until aggregates (C and D)or mature fruiting bodies (A, B, and E–G) had formed. (Scale bar, 0.5 mm.) G J tasAϪ cells have only partially lost cAR function. Using the Cre-loxP system (9), which allows recycling of the only selectable marker for Ppal, we first generated a tasBϪ mutant, and subsequently a tasAϪtasBϪ mutant. The tasBϪ mutant showed a normal phenotype (Fig. 3 A and B), but the tasAϪtasBϪ cells showed severe developmental K defects. Aggregation was normal (Fig. 3 C and D), but thereafter only small club-shaped structures were formed, consisting of thick lumpy stalks with recognizable sori (spore heads) (Fig. 3E). To confirm that this severe phenotype was due to the additional loss of TasB, we transformed the tasAϪtasBϪ cells with vector pTasBexp that contains TasB under control of its own promoter Fig. 2. Expression patterns of TasA and TasB genes. (A) Developmental (Fig. S3). The tasAϪtasBϪ/TasB mutant reverted to the pheno- regulation. Ppal cells were starved at 22 °C on water agar. Every 2 or 4 h cells Ϫ were harvested for mRNA extraction until mature fruiting bodies had formed. type of tasA cells (compare Fig. 3 F and G), indicating that the Northern blots were hybridized with 32P-dATP labeled TasA or TasB speciﬁc loss of both TasA and TasB caused the collapse of fruiting body probes. (B–J) TasB expression pattern. Ppal cells, transformed with vector morphogenesis. TasB::gal, were starved on water agar, and developing structures were ﬁxed and stained with X-Gal (37). B, aggregate; C and D, primary sorogen forma- Effects of cAR Lesions on Spore and Stalk Cell Differentiation. To test tion; E–G, seggregation of whorls; H–J, formation of secondary sorogens. whether spores and stalk cells had formed in tas null fruiting ␮ (Scale bar, 25 m.) (K) Diagram of TasB and TasA expression patterns. TasA structures, we stained their contents with Calcofluor. This dye expression was visualized earlier by expression of a TasA::GFP construct (8). fluoresces when in contact with cellulose in the walls of stalk cells and spores, but does not stain amoeboid cells. Wild-type and tasBϪ fruiting bodies form large vacuolated stalk cells in linear arrays and masses (Fig. 2 E–G). The secondary sorogens also expressed Ϫ TasB, with weaker expression at the extreme tips (Fig. 2 H–J). much smaller elliptical spores (Fig. 4 A and C). In tasA fruiting TasA was found earlier to be expressed in all cells, except those bodies, the stalk cells are more disorganized, but elliptical spores Ϫ Ϫ at the tip of the main sorogen and in the stalk (8). Therefore, the are still present (Fig. 4B). In tasA tasB fruiting bodies, disorga- expression patterns of TasB and TasA appear both to comple- nized stalk cells occupy the lower half of the structures, and no ment and partially overlap each other (Fig. 2K). elliptical spores are found. Instead, the sori contain small spherical encapsulated cells (Fig. 4D). These cells resemble the cysts or Functional Analysis of cAR Genes in Ppal. The tasAϪ cells were microcysts, which are directly formed from vegetative cells (Fig. isolated earlier from a REMI mutagenesis screen for Ppal 4F). The tasAϪtasBϪ/TasB cells again make elliptical spores and mutants with defective development. The tasAϪ fruiting bodies show the thicker stalks of tasAϪ fruiting bodies. have thick stalks and abnormal whorls, and their defective Thick Aside from their shape, spores differ from cysts by having and Aberrant Stalk gene was later identified as a cAR (8). The more condensed cytoplasm and a thick 3-layered, instead of identification of the second cAR, TasB,inPpal implies that the thinner 2-layered cell wall (10). Electron-microscopic examina-

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Fig. 5. Structural and physiological characteristics of wild-type and mutant spores and cysts. (A–C) Cell wall ultrastructure. Wild-type fruiting bodies and cysts, and tasAϪtasBϪ fruiting bodies were prepared for transmission electron microscopy. Wild-type spores are highly electron-opaque and surrounded by D a thick 3-layered cell wall (A). Wild-type cysts (B) and tasAϪtasBϪ spores (C) are less opaque with a thinner more ﬁbrous 2-layered cell wall. Overviews and procedures are listed in Fig. S4. (Scale bar, 300 nm.) (D) Resistance to lyophilization. Ppal wild-type, tasAϪtasBϪ, and tasAϪtasBϪ/TasB spores were harvested by individually lifting 4-day-old fruiting bodies from growth plates. Cysts were induced by incubating wild-type cells for 4 days in 0.3 M sorbitol. Spores and cysts were shaken for 5 min in 0.1% Triton X-100 to lyse nonen- capsulated cells, washed, and resuspended in calf serum to 104 cells/mL. One aliquot was left untreated (control), and the other lyophilized, and both samples were subsequently plated with Kaer on 3 growth plates each (diam- eter, 13 cm) at 300 cells per plate. After 4–8 days, the number of emerging colonies was counted. Data are presented as percentage of cells plated, and represent means and SD of 2 experiments.

features that are shared by tasAϪtasBϪ spores and cysts, but not by wild-type spores. Spores of all Dictyostelia can be lyophilized for storage (11), but this trait has not been demonstrated for E cysts. We measured viability after lyophilization of wild-type spores and cysts, tasAϪtasBϪ‘‘spores’’ and tasAϪtasBϪ/TasB spores. Fig. 5D shows that, in contrast to wild-type spores, cysts and tasAϪtasBϪ spores completely loose viability after lyophilization. The tasAϪtasBϪ/TasB spores are again fully resistant to lyophilization. Combined, these data show that tasAϪtasBϪ spores are actually cysts.

F Prespore Gene Induction in the tasA؊tasB؊ Mutant. We showed recently that cyst formation in Ppal only requires PKA activation (12), whereas spore formation in Ddis requires activation of both cARs and PKA (13, 14). The cues that enable PKA activation, such as SDF peptides and ammonia depletion (15), are probably still present in tasAϪtasBϪ fruiting bodies, allow- Fig. 4. Cell differentiation in Ppal cAR null mutants. (A–F) Fruiting bodies ing cysts to form. However, the absence of cARs may prevent were transferred to a droplet of 0.001% Calcoﬂuor and photographed under prespore differentiation. phase contrast (Left) and UV illumination (Right). Regular arrays of large To test this hypothesis directly, we investigated whether pre- vacuolated stalk cells and small elliptical spores are present in wild-type (A) Ϫ Ϫ Ϫ Ϫ spore differentiation is induced by cAMP in Ppal wild-type cells, and tasB (C) fruiting bodies. The tasA (B) and tasA tasB /TasB (E) fruiting Ϫ Ϫ bodies contain elliptical spores, but stalk cells are more disorganized. The but not tasA tasB mutants, using the Ppal spore coat gene SP45 lower half of tasAϪtasBϪ structures (D) consists mainly of vacuolated stalk cells (16) as a marker. Fig. 6 shows that cAMP induces SP45 Ϫ Ϫ (Bottom Left), whereas the sorus (Top Right) contains small encapsulated transcription in wild-type Ppal, but not in tasA tasB cells. These spherical cells (arrow and inset). (F) Wild-type cysts, induced by incubating data indicate that, in Ppal, cARs are essential for prespore amoebas at high osmolality (here 0.1 M KCl). (Scale bar, 5 ␮m.) induction by cAMP. Discussion Ϫ Ϫ tion of the tasA tasB spores revealed that they have the same A cAR1 Ortholog Duplicated Several Times During Dictyostelid Evo- morphology as wild-type cysts, and are dissimilar to wild-type lution. The model species Ddis has 4 cAR genes, and orthologs of spores (Fig. 5 A–C; Fig. S4). We also sought for physiological all 4 genes were found in 2 other group 4 taxa, Dmuc and Dros

7092 ͉ www.pnas.org͞cgi͞doi͞10.1073͞pnas.0901617106 Kawabe et al. Downloaded by guest on September 25, 2021 partially depend on the same signaling mechanisms. We recently gained insight in the mechanisms that control both sporulation and encystation. In Ddis, adenylate cyclase G (ACG) provides cAMP for induction of prespore differentiation in slug posteriors (21). In the fruiting body, spores are kept dormant by high osmolality, which directly activates ACG. ACG then activates PKA, resulting in inhibition of spore germination (22). In cyst forming species, such as Ppal, encystation is triggered by high osmolality, a signal for approaching drought, and this response is also mediated by ACG acting on PKA (12). Therefore, both Fig. 6. cAMP induction of prespore gene expression in Ppal cAR null mutants. Ϫ Ϫ sporulation and encystation share a requirement for PKA Ppal wild-type cells and tasA tasB mutants were incubated on agar until activation. loose aggregates had formed. Cells from dissociated loose aggregates were Although PKA activation is sufficient for encystation, sporu- shaken at 150 ϫ g and 22 °C in the absence (control) and presence of 300 ␮M cAMP, added at 60-min intervals. At 2-h intervals, cells were harvested for RNA lation additionally requires activation of cARs (23). This re- isolation. Northern blots were hybridized with a Ppal SP45 prespore DNA quirement suggests a mechanism whereby accumulation of se- probe. After stripping, the blot was rehybridized with the constitutively creted cAMP in aggregates instructs cells to form spores and not expressed Ddis 1G7 gene as a control for sample loading. cysts. It may explain why prespore induction requires much higher cAMP concentrations (Ϸ1 ␮M) (23) than chemotaxis and cAMP relay (0.1–30 nM) that also occur before aggregation (24). (Fig. 1B). The cAR genes from the earlier diverged groups 1–3 Only after aggregation can cAMP accumulate to micromolar were more similar to DdiscAR1 than to DdiscAR2-4, suggesting levels in the confined space between the cells. that cAR1 is the ancestral gene. Orthology to cAR1 was shown conclusively by the fact that the single cAR from the group 3 Scenario for the Evolution of cAMP Signaling in the Dictyostelids. species Dmin has the same flanking genes as DdiscAR1, which cAMP is a common intracellular messenger for prokaryotes and differ from those of DdiscAR2-4 (7). All tested group 1 and 3 taxa most eukaryotes, but its use as an extracellular signal is only have only a single cAR, but at least 3 independent gene well-documented for social amoebas. Intracellular cAMP has a duplications occurred in Group 2, which contains the Acytostel- deeply conserved role in sporulation and encystation of social EVOLUTION ids, the pale Polyphondylids, and 2 Dictyostelids. One gene amoebas (12), and available evidence suggests that it also duplication is evident in the Polysphondylid/Dictyostelid clade to mediates encystation of solitary amoebas (25, 26). Therefore, the form the cARs TasA and TasB. One Acytostelid, Asub, shows 2 role of extracellular cAMP is most likely derived from an duplications; in the other, Aana, only 1 cAR was detected. intracellular function, and a tentative scenario how this has Combined, these results indicate that the ancestral cAR was a occurred suggests itself. DdiscAR1 ortholog, which duplicated at least 3 times indepen- Basal dictyostelids do not use cAMP to aggregate, and at least dently during social amoeba evolution. 1 species (Dmin) uses the same attractant (folic acid) for food-seeking and for aggregation (4). Therefore, the first colo- Roles of cARs in Morphogenesis. In Ddis, propagating cAMP waves nial amoebas may have adapted their food-seeking strategy for regulate cell movement during aggregation and multicellular aggregation while still using cAMP intracellularly to trigger morphogenesis. All tested group 4 species, but none of the group encystation. Next, passive cAMP secretion and accumulation in 1–3 species use cAMP for aggregation (4). Nevertheless, phar- aggregates could have acted as a signal to prompt the starving macological experiments suggested that all Dictyostelia use cells to form spores and not cysts. Oscillatory cAMP signaling cAMP to coordinate postaggregative morphogenesis (7). We probably evolved later, first as a means to form architecturally here provide genetic evidence that fruiting body morphogenesis sophisticated fruiting structures, and finally to coordinate the crucially depends on cAR function (Fig. 3). Waves that propa- aggregation process in the most recently diverged group 4 (7). gate outward from tip regions and initiate tip-directed cell The processes that control oscillatory cAMP production by movement have been recorded in Dmin and Ddis aggregates and adenylate cyclase A share many component proteins with deeply slugs, and were identified as cAMP waves by a number of criteria conserved pathways that control the highly dynamic process of (17–20). These studies and the present observations lead to the chemotaxis in amoebas and metazoa (27–29). In the course of conclusion that primary and secondary tips on aggregates, slugs, Dictyostelid evolution, these proteins may have been recruited and whorls are autonomous cAMP oscillators that each direct a to have a novel role in the dynamic production of cAMP. group of cells to form a single unit of stalk and spore head. Our work demonstrates how a signaling pathway that mediates Therefore, oscillatory cAMP signaling is the universal and major the response of a single-celled organism to environmental stress mechanism for creating form in the social amoebas. was elaborated by adaptive evolution to coordinate developmental gene expression and morphogenetic cell movement in its Ϫ Ϫ Roles of cARs in Differentiation. The Ppal tasA tasB cells have not multicellular descendants. only lost morphogenesis, but also spore differentiation. The aggregate manages to form a short stump, which consists of Materials and Methods randomly differentiated stalk cells with a mass of cysts on top. Cell Culture. All social amoeba species were grown in association with Kleb- cARs, and therefore extracellular cAMP, appear to be essential siella aerogenes (Kaer) on LP agar or 1/5th SM agar (11). For developmental for spore differentiation. cAMP directly triggered spore gene time courses, cells were incubated at 8 ϫ 105 cells/cm2 and 22 °C on water agar expression in Ppal, and this response was lost in the tasAϪtasBϪ (1.5% agar in water) or charcoal agar (0.5% wt/vol). cells (Fig. 6). This loss is the most likely cause of its failure to form normal spores. However, unlike Ddis, where prespore cells Identification of cAR Homologs. The cAR genes were ampliﬁed by PCR from dedifferentiate in the absence of cAMP (14), in Ppal they switch genomic DNAs of 12 test species, using a mixture of primers cARdegF1/ cARdegF2 and cARdegR1/cARdegR2 (Table S1), which represent all variation to the alternative life cycle of encystation. in sequences GN/GWCWI and NPLMWR, respectively, that are conserved between cARs 1-4 of Ddis and TasA of Ppal. The PCR products were subcloned (7), Sporulation and Encystation Share Common Signaling Requirements. and their DNA sequence was determined from 3 to 20 independent clones. The The fact that spores revert to cysts in the absence of cARs almost complete 1386-bp coding sequence of the DaurcAR with 263-bp 5Ј and 208-bp certainly means that cysts are ancestral to spores, and at least 3Ј UTR was obtained by inverse PCR with primer pair DaurINV1 and DaurINV2

Kawabe et al. PNAS ͉ April 28, 2009 ͉ vol. 106 ͉ no. 17 ͉ 7093 Downloaded by guest on September 25, 2021 (Table S1), using religated ClaI digested Daur genomic (g)DNA as template. TasB promoter-LacZ construct. The TasB promoter was amplified from Ppal gDNA The 1308-bp coding region of PpalTasB with 2515-bp 5Ј UTR and 482-bp 3Ј using primers TasBpro5Ј and TasBpro3Ј, which contain XbaI and BamHI re- UTR, was obtained from 2 inverse PCR reactions using primer pairs TasBInv3/ striction sites respectively (Table S1). After digestion, the 2.3-kb PCR product TasBInv5 or TasBinvP1/TasBinvP2 (Table S1) with religated HpaI or EcoRV was ligated into BglII/XbaI digested vector pDdGal17 vector (33), yielding digested Ppal gDNAs, respectively. All PCR products were subcloned in pBlue- vector TasB::gal. scriptII (Stratagene) and sequenced. RNA isolation and analysis. Total RNA was transferred to nylon membranes (7) and hybridized at 65 °C to [32P] dATP-labeled Sp45, TasA, and TasB DNA DNA Constructs and Transformation probes. Hybridization of Ppal mRNA to a Ddis 1G7 probe was at 55 °C. Ppal LoxP-Neo vector. G418 (neomycin) resistance is the only selectable marker for TasA and TasB probes were prepared by PCR amplification of 152- and 192-bp Ppal, and has to be recycled to create double knock-out mutants. A vector with sections, respectively, of the extreme 3Ј coding region of each gene using LoxP excision sites flanking the actin6-neomycin cassette (A6neo) was con- primer pairs TasA-52/TasA-32 and TasB-55/TasB-S (Table S1) with vectors structed, which allows in vivo excision of the cassette by transient expression TasAloxP-KO and pTasBexp as templates. An 976-bp Sp45 probe was amplified of Cre-recombinase (9). The LoxP-Bsr cassette of vector pLPBLP (9) was excised from Ppal gDNA using primers Sp45F and Sp45R. with BamHI and HindIII, and ligated into pUC19 to eliminate a second XbaI site in pLPBLP. The Bsr cassette was excised with XbaI/EcoRV, leaving the LoxP sites, Phylogenetic Analysis whose flanking 5Ј overhangs were filled in with KOD DNA polymerase Alignment. Deduced amino acid sequences of partial and full-length cAR genes, (Toyobo). The actin6-NeoR cassette was excised with EcoRI/BamHI from DdisCrlA, and nondictyostelid G protein coupled receptors (GPCRs) with the pB10SX (30), filled in, and blunt-end ligated into pUC19-LoxP, yielding vector pfam05462 Dicty࿝CAR domain (34) were aligned with CLUSTALW (35), pLoxNeoI. using the region spanning the first 6 transmembrane helices. The cARs Vectors for TasA and TasB gene disruption. Partial TasB sequence with 0.8-kb 5Ј aligned unambiguously, but the alignment of DdisCrlA and nondictyos- UTR and 2.2-kb 3Ј UTR was amplified by inverse PCR from HpaI digested and telid GPCRs was edited according to their individual alignment to the religated Ppal gDNA, using primers TasBinvK1 and TasBinvK2 (Table S1). The pfam05462 domain. BamHI digested PCR product was cloned into BamHI digested pLoxNeoI Phylogeny. A cAR phylogeny was constructed by Bayesian inference (36). A yielding TasBloxP-KO. To obtain a TasA disruption vector, partial TasA se- mixed amino acid model was used with rate variation across sites estimated by Ј Ј quence with 1.4-kb 5 UTR and 1.1-kb 3 UTR was amplified with primers a gamma distribution with 6 rate categories and no invariable sites. The TasAinvK1 and TasAinvK2 from XbaI digested and religated Ppal gDNA, and analysis was run for 106 generations, at which point the average SD of split cloned as above in pLoxNeoI, yielding TasAloxP-KO. Cells were transformed as frequencies was 0.0074. Posterior probabilities were averaged over the final described previously (31), and screened for homologous recombination by 2 75% of trees. The analysis was also run with other prior settings such as separate PCR reactions and analysis of Southern blots (for detailed proce- introduction of a proportion of invariable sites, reducing rate categories to 4, dures, see Fig. S5 and Fig. S6). To remove the A6neo cassette, knock-out cells or setting rate variation to equal. These changes did not affect tree topology. were transformed with pA15NLS.Cre for transient expression of Cre- recombinase. Transformed clones were replica-plated onto autoclaved Kaer ACKNOWLEDGMENTS. We thank Drs. Jan Faix (Ludwig Maximilians Univer- ␮ on LP agar with and without 200 g/mL G418 for negative selection. sity, Munich) and Lisa Kreppel and Alan Kimmel (National Institutes of Health, TasB expression vector. To express TasB from its own promoter, the TasB coding Bethesda, MD) for their kind gifts of vectors pLPBLP and pA15NLS.cre. This sequence and promoter were amplified separately by PCR, sequentially cloned work was supported by Biotechnology and Biological Sciences Research Coun- into vector pEXP5 (32), and introduced into tasAϪtasBϪneoϪ cells (Fig. S3). cil Grant BB/D013453/1 and Wellcome Trust Grant 076618.

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