• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of pnasPNASInfo for AuthorsSubscriptionsAboutThis Article
Proc Natl Acad Sci U S A. Apr 28, 2009; 106(17): 7089–7094.
Published online Apr 14, 2009. doi:  10.1073/pnas.0901617106
PMCID: PMC2678454
Evolution

Activated cAMP receptors switch encystation into sporulation

Abstract

Metazoan embryogenesis is controlled by a limited number of signaling modules that are used repetitively at successive developmental stages. The development of social amoebas shows similar reiterated use of cAMP-mediated signaling. In the model Dictyostelium discoideum, secreted cAMP acting on 4 cAMP receptors (cARs1-4) coordinates cell movement during aggregation and fruiting body formation, and induces the expression of aggregation and sporulation genes at consecutive developmental stages. To identify hierarchy in the multiple roles of cAMP, we investigated cAR heterogeneity and function across the social amoeba phylogeny. The gene duplications that yielded cARs 2-4 occurred late in evolution. Many species have only a cAR1 ortholog that duplicated independently in the Polysphondylids and Acytostelids. Disruption of both cAR genes of Polysphondylium pallidum (Ppal) did not affect aggregation, but caused complete collapse of fruiting body morphogenesis. The stunted structures contained disorganized stalk cells, which supported a mass of cysts instead of spores; cAMP triggered spore gene expression in Ppal, but not in the cAR null mutant, explaining its sporulation defect. Encystation is the survival strategy of solitary amoebas, and lower taxa, like Ppal, can still encyst as single cells. Recent findings showed that intracellular cAMP accumulation suffices to trigger encystation, whereas it is a complementary requirement for sporulation. Combined, the data suggest that cAMP signaling in social amoebas evolved from cAMP-mediated encystation in solitary amoebas; cAMP secretion in aggregates prompted the starving cells to form spores and not cysts, and additionally organized fruiting body morphogenesis. cAMP-mediated aggregation was the most recent innovation.

Keywords: developmental signaling, evolution of multicellularity, Dictyostelia, Amoebozoa

The evolution of novel morphological features is due to changes in the developmental signaling processes that shape these features. The number of different signals that shape complex embryos, such as mammals, is surprisingly limited because the same signals, such as members of the wingless/wnt, hedgehog, TGF-β, and FGF families, are used many times over at successive developmental stages (13). Also, the signals and their associated transduction pathways are deeply conserved in evolution, often having homologous roles in shaping the body plan of lower invertebrates. The first multicellular organisms most likely deployed preexisting signaling systems from their unicellular ancestors, which were used to find food or mates, or to evade stress. To fully understand developmental signaling, it is of fundamental importance to identify which protist signaling pathways were used, and how they were adapted and elaborated to generate the ever increasing complexity of multicellular organisms.

The dictyostelid social amoebas offer unique opportunities to address this issue. They are as genetically diverse as animals, but alternate a sophisticated program of multicellular morphogenesis with a free-living amoeboid lifestyle. A robust molecular phylogeny is available, showing subdivision of all known species into 4 groups, with the model Dictyostelium discoideum residing in the most derived group 4 (4). During D. discoideum development, the deeply conserved intracellular messenger cAMP has multiple roles as a secreted signal, detected by 4 homologous cAMP receptors (cAR1–4) (5). cAMP pulses coordinate the aggregation of starving cells and organize the construction of fruiting bodies with a highly regulated pattern of spores and stalk cells. Secreted cAMP also up-regulates expression of aggregation genes, induces expression of spore genes, and inhibits stalk gene expression (6).

Single cAR genes were previously detected in 3 more basal dictyostelid taxa, but were only expressed after aggregation. The nonhydrolyzable cAMP analog SpcAMPS, which inhibits cAR-mediated pulsatile cAMP signaling, disorganized fruiting body formation in these taxa, suggesting an ancestral role for pulsatile cAMP signaling in fruiting body morphogenesis (7). However, it remains unresolved whether cARs also mediated gene expression in basal taxa, for which SpcAMPS is a normal agonist, and whether the detected cARs were unique or members of larger gene families.

To resolve these questions, we first mapped cAR heterogeneity in the Dictyostelia, showing multiple independent events of cAR gene duplication. Second, by successively disrupting all cAR genes of the early diverged taxon Polysphondylium pallidum (Ppal), we greatly expanded the opportunities for cAR functional analysis. Loss of 1 cAR caused reduced fruiting body branching, whereas loss of both disrupted fruiting body formation. Strikingly, without cARs, Ppal cells could not express spore genes, and formed cysts instead of spores in the stunted structures. Encystation is the major stress response of solitary protists, which is retained in several early diverging social amoebas. Our data show that cysts are ancestral to spores, and that activated cARs determine the choice between the 2 developmental pathways.

Results

cAR Heterogeneity in the Dictyostelia.

D. discoideum has 4 homologous cARs with different functions in chemotaxis and gene regulation (5). To understand how these functions evolved, we first mapped patterns of cAR gene duplication across the Dictyostelid phylogeny. Two to 5 test species were selected from each of the 4 groups of Dictyostelia (Fig. 1A). Amplification of cAR genes from genomic DNAs of the selected species yielded 1 or multiple products ranging from 324 to 330 bp. Their derived amino acid sequences shared 59–97% sequence identity between each other and DdiscARs (Fig. S1A). Four cAR homologs each, were amplified from the group 4 species Dictyostelium mucoroides (Dmuc) and Dictyostelium rosarium (Dros). Ppal and 2 related Group 2 species, excluding the Acytostelids, yielded 2 cAR homologs each. Acytostelium subglobosum (Asub) yielded 3 cAR homologs, but only a single cAR was isolated from Acytostelium anastomosans (Aana). All tested Group 1 and 3 species, and the group-intermediate species Polysphondylium violaceum (Pvio) contained only a single cAR homolog.

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 ...

We next isolated full-length cAR genes by inverse PCR from at least 1 species in each group. Full-length sequences were already available for DdiscARs 1-4 in group 4 (5), Dictyostelium minutum (Dmin) cAR in group 3 (7), and 1 Ppal cAR, named TasA in group 2 (8). Complete coding regions were amplified for the Dictyostelium aureo-stipes (Daur) cAR in group 1, and for the second cAR of Ppal, which was named TasB. Alignment of the deduced amino acid sequences of DaurcAR and PpalTasB with other complete cARs shows that the regions that contain the 7 transmembrane helices are highly conserved, whereas the carboxyl-termini of the cARs are very diverse (Fig. S1B).

An alignment of all PCR products and full-length cARs was used to assess phylogenetic relationships between the cARs. Fig. 1B shows that the 4 cARs of Dmuc and Dros are most similar to cARs1-4 in Ddis. The 2 cARs that were identified in Dictyostelium gloeosporum (Dglo) and Polysphondylium pseudocandidum (Ppse), 2 species that are closely related to Ppal (Fig. 1A), are most similar Ppal TasA or TasB. The 3 Asub cARs are quite diverse, but still more similar to cARs than to the Ddis cAR-like protein CrlA. All nongroup 4 cARs are more similar to DdiscAR1 than to DdiscARs2-4. These data indicate that the gene duplications that gave rise to cARs2-4 occurred in Group 4 taxa, and that independent cAR gene duplications occurred at least 3 more times.

Expression Patterns of cARs in Ppal.

We planned to use the large genomic fragments of the Dmin and DaurcARs and PpalTasB to generate constructs for expression analysis and gene disruption. However, we have not been able to achieve stable transformation of Dmin and Daur cells yet. Ppal can be transformed; therefore, we concentrated on analyzing cAR function in this species. We first compared the temporal expression patterns of TasB and TasA. Fig. 2A shows that TasB expression is first visible after 2 h of starvation, to reach a plateau at 8 h when aggregation is completed. TasA mRNA first appears after aggregates have formed. Both genes remain highly expressed until late fruiting body formation. To visualize the spatial pattern of TasB expression, its 2.3-kb promoter was fused to the LacZ reporter gene and transformed into Ppal cells. Developing structures were incubated with X-Gal to visualize β-galactosidase activity. TasB was first expressed weakly at the center of streaming aggregates (Fig. 2B). Expression then increased throughout the newly formed sorogen, but soon appeared strongest at the tip region (Fig. 2 C and D). Ppal fruiting bodies form whorls of side branches from cell masses that pinch off from the rear of the sorogen. TasB showed high expression at the site of separation of these cell masses (Fig. 2 E–G). The secondary sorogens also expressed TasB, with weaker expression at the extreme tips (Fig. 2 H–J). TasA was found earlier to be expressed in all cells, except those at the tip of the main sorogen and in the stalk (8). Therefore, the expression patterns of TasB and TasA appear both to complement and partially overlap each other (Fig. 2K).

Fig. 2.
Expression patterns of TasA and TasB genes. (A) Developmental regulation. Ppal cells were starved at 22 °C on water agar. Every 2 or 4 h cells were harvested for mRNA extraction until mature fruiting bodies had formed. Northern blots were hybridized ...

Functional Analysis of cAR Genes in Ppal.

The tasA cells were isolated earlier from a REMI mutagenesis screen for Ppal mutants with defective development. The tasA fruiting bodies have thick stalks and abnormal whorls, and their defective Thick and Aberrant Stalk gene was later identified as a cAR (8). The identification of the second cAR, TasB, in Ppal implies that the tasA 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 tasB mutant, and subsequently a tasAtasB mutant.

The tasB mutant showed a normal phenotype (Fig. 3 A and B), but the tasAtasB cells showed severe developmental 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 tasAtasB cells with vector pTasBexp that contains TasB under control of its own promoter (Fig. S3). The tasAtasB/TasB mutant reverted to the phenotype of tasA cells (compare Fig. 3 F and G), indicating that the loss of both TasA and TasB caused the collapse of fruiting body morphogenesis.

Fig. 3.
Phenotypes of single and double tasA and tasB mutants. The Ppal tasA mutant was created earlier (8), and the generation of tasB, tasAtasB, and tasAtasB/TasB mutants is outlined in ...

Effects of cAR Lesions on Spore and Stalk Cell Differentiation.

To test whether spores and stalk cells had formed in tas null fruiting structures, we stained their contents with Calcofluor. This dye fluoresces when in contact with cellulose in the walls of stalk cells and spores, but does not stain amoeboid cells. Wild-type and tasB fruiting bodies form large vacuolated stalk cells in linear arrays and much smaller elliptical spores (Fig. 4 A and C). In tasA fruiting bodies, the stalk cells are more disorganized, but elliptical spores are still present (Fig. 4B). In tasAtasB fruiting bodies, disorganized stalk cells occupy the lower half of the structures, and no elliptical spores are found. Instead, the sori contain small spherical encapsulated cells (Fig. 4D). These cells resemble the cysts or microcysts, which are directly formed from vegetative cells (Fig. 4F). The tasAtasB/TasB cells again make elliptical spores and show the thicker stalks of tasA fruiting bodies.

Fig. 4.
Cell differentiation in Ppal cAR null mutants. (A–F) Fruiting bodies were transferred to a droplet of 0.001% Calcofluor and photographed under phase contrast (Left) and UV illumination (Right). Regular arrays of large vacuolated stalk cells and ...

Aside from their shape, spores differ from cysts by having more condensed cytoplasm and a thick 3-layered, instead of thinner 2-layered cell wall (10). Electron-microscopic examination of the tasAtasB spores revealed that they have the same morphology as wild-type cysts, and are dissimilar to wild-type spores (Fig. 5 A–C; Fig. S4). We also sought for physiological features that are shared by tasAtasB 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 cysts. We measured viability after lyophilization of wild-type spores and cysts, tasAtasB“spores” and tasAtasB/TasB spores. Fig. 5D shows that, in contrast to wild-type spores, cysts and tasAtasB spores completely loose viability after lyophilization. The tasAtasB/TasB spores are again fully resistant to lyophilization. Combined, these data show that tasAtasB spores are actually cysts.

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 tasAtasB fruiting bodies were prepared for transmission electron ...

Prespore Gene Induction in the tasAtasB 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 tasAtasB fruiting bodies, allowing cysts to form. However, the absence of cARs may prevent prespore differentiation.

To test this hypothesis directly, we investigated whether prespore differentiation is induced by cAMP in Ppal wild-type cells, but not tasAtasB mutants, using the Ppal spore coat gene SP45 (16) as a marker. Fig. 6 shows that cAMP induces SP45 transcription in wild-type Ppal, but not in tasAtasB cells. These data indicate that, in Ppal, cARs are essential for prespore induction by cAMP.

Fig. 6.
cAMP induction of prespore gene expression in Ppal cAR null mutants. Ppal wild-type cells and tasAtasB mutants were incubated on agar until loose aggregates had formed. Cells from dissociated loose aggregates were shaken at 150 × ...

Discussion

A cAR1 Ortholog Duplicated Several Times During Dictyostelid Evolution.

The model species Ddis has 4 cAR genes, and orthologs of all 4 genes were found in 2 other group 4 taxa, Dmuc and Dros (Fig. 1B). The cAR genes from the earlier diverged groups 1–3 were more similar to DdiscAR1 than to DdiscAR2-4, suggesting that cAR1 is the ancestral gene. Orthology to cAR1 was shown conclusively by the fact that the single cAR from the group 3 species Dmin has the same flanking genes as DdiscAR1, which differ from those of DdiscAR2-4 (7). All tested group 1 and 3 taxa have only a single cAR, but at least 3 independent gene duplications occurred in Group 2, which contains the Acytostelids, the pale Polyphondylids, and 2 Dictyostelids. One gene duplication is evident in the Polysphondylid/Dictyostelid clade to form the cARs TasA and TasB. One Acytostelid, Asub, shows 2 duplications; in the other, Aana, only 1 cAR was detected. Combined, these results indicate that the ancestral cAR was a DdiscAR1 ortholog, which duplicated at least 3 times independently during social amoeba evolution.

Roles of cARs in Morphogenesis.

In Ddis, propagating cAMP waves regulate cell movement during aggregation and multicellular morphogenesis. All tested group 4 species, but none of the group 1–3 species use cAMP for aggregation (4). Nevertheless, pharmacological experiments suggested that all Dictyostelia use cAMP to coordinate postaggregative morphogenesis (7). We here provide genetic evidence that fruiting body morphogenesis crucially depends on cAR function (Fig. 3). Waves that propagate outward from tip regions and initiate tip-directed cell movement have been recorded in Dmin and Ddis aggregates and slugs, and were identified as cAMP waves by a number of criteria (1720). These studies and the present observations lead to the conclusion that primary and secondary tips on aggregates, slugs, and whorls are autonomous cAMP oscillators that each direct a group of cells to form a single unit of stalk and spore head. Therefore, oscillatory cAMP signaling is the universal and major mechanism for creating form in the social amoebas.

Roles of cARs in Differentiation.

The Ppal tasAtasB cells have not only lost morphogenesis, but also spore differentiation. The aggregate manages to form a short stump, which consists of randomly differentiated stalk cells with a mass of cysts on top. cARs, and therefore extracellular cAMP, appear to be essential for spore differentiation. cAMP directly triggered spore gene expression in Ppal, and this response was lost in the tasAtasB cells (Fig. 6). This loss is the most likely cause of its failure to form normal spores. However, unlike Ddis, where prespore cells dedifferentiate in the absence of cAMP (14), in Ppal they switch to the alternative life cycle of encystation.

Sporulation and Encystation Share Common Signaling Requirements.

The fact that spores revert to cysts in the absence of cARs almost certainly means that cysts are ancestral to spores, and at least 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 sporulation and encystation share a requirement for PKA activation.

Although PKA activation is sufficient for encystation, sporulation additionally requires activation of cARs (23). This requirement suggests a mechanism whereby accumulation of secreted cAMP in aggregates instructs cells to form spores and not cysts. It may explain why prespore induction requires much higher cAMP concentrations (≈1 μM) (23) than chemotaxis and cAMP relay (0.1–30 nM) that also occur before aggregation (24). Only after aggregation can cAMP accumulate to micromolar levels in the confined space between the cells.

Scenario for the Evolution of cAMP Signaling in the Dictyostelids.

cAMP is a common intracellular messenger for prokaryotes and most eukaryotes, but its use as an extracellular signal is only well-documented for social amoebas. Intracellular cAMP has a deeply conserved role in sporulation and encystation of social amoebas (12), and available evidence suggests that it also mediates encystation of solitary amoebas (25, 26). Therefore, the role of extracellular cAMP is most likely derived from an intracellular function, and a tentative scenario how this has occurred suggests itself.

Basal dictyostelids do not use cAMP to aggregate, and at least 1 species (Dmin) uses the same attractant (folic acid) for food-seeking and for aggregation (4). Therefore, the first colonial amoebas may have adapted their food-seeking strategy for aggregation while still using cAMP intracellularly to trigger encystation. Next, passive cAMP secretion and accumulation in aggregates could have acted as a signal to prompt the starving cells to form spores and not cysts. Oscillatory cAMP signaling probably evolved later, first as a means to form architecturally sophisticated fruiting structures, and finally to coordinate the aggregation process in the most recently diverged group 4 (7). The processes that control oscillatory cAMP production by adenylate cyclase A share many component proteins with deeply conserved pathways that control the highly dynamic process of chemotaxis in amoebas and metazoa (2729). In the course of Dictyostelid evolution, these proteins may have been recruited to have a novel role in the dynamic production of cAMP.

Our work demonstrates how a signaling pathway that mediates the response of a single-celled organism to environmental stress was elaborated by adaptive evolution to coordinate developmental gene expression and morphogenetic cell movement in its multicellular descendants.

Materials and Methods

Cell Culture.

All social amoeba species were grown in association with Klebsiella aerogenes (Kaer) on LP agar or 1/5th SM agar (11). For developmental time courses, cells were incubated at 8 × 105 cells/cm2 and 22 °C on water agar (1.5% agar in water) or charcoal agar (0.5% wt/vol).

Identification of cAR Homologs.

The cAR genes were amplified by PCR from genomic DNAs of 12 test species, using a mixture of primers cARdegF1/cARdegF2 and cARdegR1/cARdegR2 (Table S1), which represent all variation 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), and their DNA sequence was determined from 3 to 20 independent clones. The complete 1386-bp coding sequence of the DaurcAR with 263-bp 5′ and 208-bp 3′ UTR was obtained by inverse PCR with primer pair DaurINV1 and DaurINV2 (Table S1), using religated ClaI digested Daur genomic (g)DNA as template. The 1308-bp coding region of PpalTasB with 2515-bp 5′ UTR and 482-bp 3′ UTR, was obtained from 2 inverse PCR reactions using primer pairs TasBInv3/TasBInv5 or TasBinvP1/TasBinvP2 (Table S1) with religated HpaI or EcoRV digested Ppal gDNAs, respectively. All PCR products were subcloned in pBluescriptII (Stratagene) and sequenced.

DNA Constructs and Transformation

LoxP-Neo vector.

G418 (neomycin) resistance is the only selectable marker for Ppal, and has to be recycled to create double knock-out mutants. A vector with LoxP excision sites flanking the actin6-neomycin cassette (A6neo) was constructed, which allows in vivo excision of the cassette by transient expression of Cre-recombinase (9). The LoxP-Bsr cassette of vector pLPBLP (9) was excised 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, whose flanking 5′ overhangs were filled in with KOD DNA polymerase (Toyobo). The actin6-NeoR cassette was excised with EcoRI/BamHI from pB10SX (30), filled in, and blunt-end ligated into pUC19-LoxP, yielding vector pLoxNeoI.

Vectors for TasA and TasB gene disruption.

Partial TasB sequence with 0.8-kb 5′ UTR and 2.2-kb 3′ UTR was amplified by inverse PCR from HpaI digested and religated Ppal gDNA, using primers TasBinvK1 and TasBinvK2 (Table S1). The BamHI digested PCR product was cloned into BamHI digested pLoxNeoI yielding TasBloxP-KO. To obtain a TasA disruption vector, partial TasA sequence with 1.4-kb 5′ UTR and 1.1-kb 3′ UTR was amplified with primers TasAinvK1 and TasAinvK2 from XbaI digested and religated Ppal gDNA, and cloned as above in pLoxNeoI, yielding TasAloxP-KO. Cells were transformed as described previously (31), and screened for homologous recombination by 2 separate PCR reactions and analysis of Southern blots (for detailed procedures, see Fig. S5 and Fig. S6). To remove the A6neo cassette, knock-out cells were transformed with pA15NLS.Cre for transient expression of Cre-recombinase. Transformed clones were replica-plated onto autoclaved Kaer on LP agar with and without 200 μg/mL G418 for negative selection.

TasB expression vector.

To express TasB from its own promoter, the TasB coding sequence and promoter were amplified separately by PCR, sequentially cloned into vector pEXP5 (32), and introduced into tasAtasBneo cells (Fig. S3).

TasB promoter-LacZ construct.

The TasB promoter was amplified from Ppal gDNA using primers TasBpro5′ and TasBpro3′, which contain XbaI and BamHI restriction sites respectively (Table S1). After digestion, the 2.3-kb PCR product was ligated into BglII/XbaI digested vector pDdGal17 vector (33), yielding vector TasB::gal.

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 probes. Hybridization of Ppal mRNA to a Ddis 1G7 probe was at 55 °C. Ppal TasA and TasB probes were prepared by PCR amplification of 152- and 192-bp sections, respectively, of the extreme 3′ coding region of each gene using primer pairs TasA-52/TasA-32 and TasB-55/TasB-S (Table S1) with vectors TasAloxP-KO and pTasBexp as templates. An 976-bp Sp45 probe was amplified from Ppal gDNA using primers Sp45F and Sp45R.

Phylogenetic Analysis

Alignment.

Deduced amino acid sequences of partial and full-length cAR genes, DdisCrlA, and nondictyostelid G protein coupled receptors (GPCRs) with the pfam05462 Dicty_CAR domain (34) were aligned with CLUSTALW (35), using the region spanning the first 6 transmembrane helices. The cARs aligned unambiguously, but the alignment of DdisCrlA and nondictyostelid GPCRs was edited according to their individual alignment to the pfam05462 domain.

Phylogeny.

A cAR phylogeny was constructed by Bayesian inference (36). A mixed amino acid model was used with rate variation across sites estimated by a gamma distribution with 6 rate categories and no invariable sites. The analysis was run for 106 generations, at which point the average SD of split frequencies was 0.0074. Posterior probabilities were averaged over the final 75% of trees. The analysis was also run with other prior settings such as introduction of a proportion of invariable sites, reducing rate categories to 4, or setting rate variation to equal. These changes did not affect tree topology.

Supplementary Material

Supporting Information:

Acknowledgments.

We thank Drs. Jan Faix (Ludwig Maximilians University, Munich) and Lisa Kreppel and Alan Kimmel (National Institutes of Health, Bethesda, MD) for their kind gifts of vectors pLPBLP and pA15NLS.cre. This work was supported by Biotechnology and Biological Sciences Research Council Grant BB/D013453/1 and Wellcome Trust Grant 076618.

Footnotes

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Data deposition: The sequences reported in this paper have been deposited in the GenBank database [accession nos. EU797651 (DroscAR3), EU797652 (DroscAR4), EU797653 (DmuccAR1), EU797654 (DmuccAR2), EU797655 (DmuccAR3), EU797656 (DmuccAR4), EU797657 (PviocAR), EU797658 (DrhicAR), EU797668 (PpalTasB), EU797661 (PpseTasA), EU797662 (PpseTasB), EU797663 (DgloTasA), EU797664 (DgloTasB), EU797660 (AanacAR), EU797665 (AsubcARE), EU797666 (AsubcARF), EU797667 (AsubcARG), EU797659 (DbifcAR), and EU797669 (DaurcAR)].

This article contains supporting information online at www.pnas.org/cgi/content/full/0901617106/DCSupplemental.

References

1. True JR, Carroll SB. Gene co-option in physiological and morphological evolution. Annu Rev Cell Dev Biol. 2002;18:53–80. [PubMed]
2. Guder C, et al. The Wnt code: Cnidarians signal the way. Oncogene. 2006;25:7450–7460. [PubMed]
3. Varjosalo M, Taipale J. Hedgehog: Functions and mechanisms. Gene Dev. 2008;22:2454–2472. [PubMed]
4. Schaap P, et al. Molecular phylogeny and evolution of morphology in the social amoebas. Science. 2006;314:661–663. [PMC free article] [PubMed]
5. Parent CA, Devreotes PN. Molecular genetics of signal transduction in Dictyostelium. Annu Rev Biochem. 1996;65:411–440. [PubMed]
6. Aubry L, Firtel R. Integration of signaling networks that regulate Dictyostelium differentiation. Ann Rev Cell Dev Biol. 1999;15:469–517. [PubMed]
7. Alvarez-Curto E, et al. Evolutionary origin of cAMP-based chemoattraction in the social amoebae. Proc Natl Acad Sci USA. 2005;102:6385–6390. [PMC free article] [PubMed]
8. Kawabe Y, Kuwayama H, Morio T, Urushihara H, Tanaka Y. A putative serpentine receptor gene tasA required for normal morphogenesis of primary stalk and branch structure in Polysphondylium pallidum. Gene. 2002;285:291–299. [PubMed]
9. Faix J, Kreppel L, Shaulsky G, Schleicher M, Kimmel AR. A rapid and efficient method to generate multiple gene disruptions in Dictyostelium discoideum using a single selectable marker and the Cre-loxP system. Nucleic Acids Res. 2004;32:e143. [PMC free article] [PubMed]
10. Hohl HR, Miura-Santo LY, Cotter DA. Ultrastuctural changes during formation and germination of microcysts in Polysphondylium pallidum, a cellular slime mould. J Cell Sci. 1970;7:285–306. [PubMed]
11. Raper KB. The Dictyostelids. Princeton: Princeton Univ Press; 1984.
12. Ritchie AV, van Es S, Fouquet C, Schaap P. From drought sensing to developmental control: Evolution of cyclic AMP signaling in social amoebas. Mol Biol Evol. 2008;25:2109–2118. [PMC free article] [PubMed]
13. Hopper NA, Harwood AJ, Bouzid S, Véron M, Williams JG. Activation of the prespore and spore cell pathway of Dictyostelium differentiation by cAMP-dependent protein kinase and evidence for its upstream regulation by ammonia. EMBO J. 1993;12:2459–2466. [PMC free article] [PubMed]
14. Wang M, Van Driel R, Schaap P. Cyclic AMP-phosphodiesterase induces dedifferentiation of prespore cells in Dictyostelium discoideum slugs: Evidence that cyclic AMP is the morphogenetic signal for prespore differentiation. Development. 1988;103:611–618.
15. Anjard C, Zeng CJ, Loomis WF, Nellen W. Signal transduction pathways leading to spore differentiation in Dictyostelium discoideum. Dev Biol. 1998;193:146–155. [PubMed]
16. Gregg KY, Cox EC. Spatial and temporal expression of a Polysphondylium spore-specific gene. Dev Biol. 2000;224:81–95. [PubMed]
17. Schaap P, Konijn TM, Van Haastert PJM. cAMP pulses coordinate morphogenetic movement during fruiting body formation of Dictyostelium minutum. Proc Natl Acad Sci USA. 1984;81:2122–2126. [PMC free article] [PubMed]
18. Schaap P. cAMP relay during early culmination of Dictyostelium minutum. Differentiation. 1985;28:205–208. [PubMed]
19. Siegert F, Weijer CJ. Three-dimensional scroll waves organize Dictyostelium slugs. Proc Natl Acad Sci USA. 1992;89:6433–6437. [PMC free article] [PubMed]
20. Dormann D, Abe T, Weijer CJ, Williams J. Inducible nuclear translocation of a STAT protein in Dictyostelium prespore cells: Implications for morphogenesis and cell-type regulation. Development. 2001;128:1081–1088. [PubMed]
21. Alvarez-Curto E, et al. cAMP production by adenylyl cyclase G induces prespore differentiation in Dictyostelium slugs. Development. 2007;134:959–966. [PMC free article] [PubMed]
22. Van Es S, et al. Adenylyl cyclase G, an osmosensor controlling germination of Dictyostelium spores. J Biol Chem. 1996;271:23623–23625. [PubMed]
23. Schaap P, Van Driel R. Induction of post-aggregative differentiation in Dictyostelium discoideum by cAMP. Evidence of involvement of the cell surface cAMP receptor. Exp Cell Res. 1985;159:388–398. [PubMed]
24. Van Haastert PJM, Konijn TM. Signal transduction in the cellular slime molds. Mol Cell Endocrinol. 1982;26:1–17. [PubMed]
25. Raizada MK, Murti CRK. Transformation of trophic Hartmannella culbertsoni into viable cysts by cyclic 3′5′-adenosine monophosphate. J Cell Biol. 1972;52:743–748. [PMC free article] [PubMed]
26. Coppi A, Merali S, Eichinger D. The enteric parasite Entamoeba uses an autocrine catecholamine system during differentiation into the infectious cyst stage. J Biol Chem. 2002;277:8083–8090. [PubMed]
27. Lilly PJ, Devreotes PN. Chemoattractant and GTPgS-mediated stimulation of adenylyl cyclase in Dictyostelium requires translocation of CRAC to membranes. J Cell Biol. 1995;129:1659–1665. [PMC free article] [PubMed]
28. Comer FI, Lippincott CK, Masbad JJ, Parent CA. The PI3K-mediated activation of CRAC independently regulates adenylyl cyclase activation and chemotaxis. Curr Biol. 2005;15:134–139. [PubMed]
29. Lee S, et al. TOR complex 2 integrates cell movement during chemotaxis and signal relay in Dictyostelium. Mol Biol Cell. 2005;16:4572–4583. [PMC free article] [PubMed]
30. Nellen W, Firtel RA. High-copy-number transformants and co-transformation in Dictyostelium. Gene. 1985;39:155–163. [PubMed]
31. Kawabe Y, Enomoto T, Morio T, Urushihara H, Tanaka Y. LbrA, a protein predicted to have a role in vesicle trafficking, is necessary for normal morphogenesis in Polysphondylium pallidum. Gene. 1999;239:75–79. [PubMed]
32. Meima ME, Weening KE, Schaap P. Vectors for expression of proteins with single or combinatorial fluorescent protein and tandem affinity purification tags in Dictyostelium. Protein Expr Purif. 2007;53:283–288. [PMC free article] [PubMed]
33. Harwood AJ, Drury L. New vectors for expression of the E. coli lacZ gene in Dictyostelium. Nucleic Acids Res. 1990;18:4292. [PMC free article] [PubMed]
34. Bateman A, et al. The Pfam protein families database. Nucleic Acids Res. 2004;32:D138–D141. [PMC free article] [PubMed]
35. Chenna R, et al. Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res. 2003;31:3497–3500. [PMC free article] [PubMed]
36. Ronquist F, Huelsenbeck JP. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics. 2003;19:1572–1574. [PubMed]
37. Dingermann T, et al. Optimization and in situ detection of Escherichia coli beta-galactosidase gene expression in Dictyostelium discoideum. Gene. 1989;85:353–362. [PubMed]

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...