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. 2005 Dec;171(4):1523–1533. doi: 10.1534/genetics.105.047233

Schizosaccharomyces pombe Adenylate Cyclase Suppressor Mutations Suggest a Role for cAMP Phosphodiesterase Regulation in Feedback Control of Glucose/cAMP Signaling

Lili Wang 1, Kenneth Griffiths Jr 1,1, Y Hi Zhang 1, F Douglas Ivey 1, Charles S Hoffman 1,2
PMCID: PMC1456081  PMID: 16143612

Abstract

Mutations affecting the Schizosaccharomyces pombe cAMP phosphodiesterase (PDE) gene cgs2+ were identified in a screen for suppressors of mutant alleles of the adenylate cyclase gene (git2+/cyr1+), which encode catalytically active forms of the enzyme that cannot be stimulated by extracellular glucose signaling. These mutations suppress both the git2− mutant alleles used in the suppressor selection and mutations in git1+, git3+, git5+, git7+, git10+, and git11+, which are all required for adenylate cyclase activation. Notably, these cgs2 mutant alleles fail to suppress mutations in gpa2+, which encodes the Gα subunit of a heterotrimeric G protein required for adenylate cyclase activation, although the previously identified cgs2-2 allele does suppress loss of gpa2+. Further analysis of the cgs2-s1 allele reveals a synthetic interaction with the gpa2R176H-activated allele, with respect to derepression of fbp1-lacZ transcription in glucose-starved cells. In addition, direct measurements of cAMP levels show that cgs2-s1 cells maintain normal basal cAMP levels, but are severely defective in feedback regulation upon glucose detection. These results suggest that PDE activity in S. pombe may be coordinately regulated with adenylate cyclase activity as part of the feedback regulation mechanism to limit the cAMP response to glucose detection.

NUTRIENT sensing is a critical process for microorganisms as they must regulate their growth and metabolism in response to changes in the growth environment. Signal transduction pathways that are activated by either the presence or the absence of specific nutrients, the best studied of which is glucose, have been identified. Both yeast and fungi, including a wide variety of pathogenic fungi, have been shown to regulate metabolic pathways, sexual development, and growth morphology in response to nutrient signaling (Lengeler et al. 2000).

In the budding yeast Saccharomyces cerevisiae and the fission yeast Schizosaccharomyces pombe, glucose detection leads to a transient cAMP signal, which activates the cAMP-dependent protein kinase PKA. The increase in cAMP is due to the activation of adenylate cyclase, which converts ATP to cAMP, and not to the inactivation of cAMP phosphodiesterase (PDE), which converts cAMP to AMP (Nikawa et al. 1987; Byrne and Hoffman 1993). Genetic and molecular analyses to characterize the mechanism of glucose detection and adenylate cyclase activation have uncovered some related components between these two cAMP signaling pathways, as well as several features that are unique to each pathway (Hoffman 2005). For example, both yeasts express seven-transmembrane G-protein-coupled receptors (GPCRs; S. cerevisiae Gpr1 and S. pombe Git3), which are required to activate similar Gpa2 Gα subunits (Nocero et al. 1994; Xue et al. 1998; Yun et al. 1998; Kraakman et al. 1999; Lorenz et al. 2000; Welton and Hoffman 2000). However, S. pombe Gpa2, which has recently been shown to directly bind an N-terminal domain of adenylate cyclase (Ivey and Hoffman 2005), functions together with the Git5-Git11 Gβγ dimer (Landry et al. 2000; Landry and Hoffman 2001), while the only authentic S. cerevisiae Gβγ dimer, Ste4-Ste18, does not act in conjunction with Gpa2 (Liu et al. 1993). In addition, Ras proteins play an important role in adenylate cyclase activation in S. cerevisiae, but have no role in the S. pombe cAMP pathway (Toda et al. 1985; Fukui et al. 1986; Mbonyi et al. 1988; Hoffman and Winston 1991).

The mechanism of fungal adenylate cyclase activation remains largely unknown. In budding yeast, the GTP-binding Ras proteins have been shown to bind to a sequence far from the catalytic domain, while, in mammals, G proteins bind directly to adenylate cyclase catalytic domains. In S. pombe, most of the genes known to encode components of the glucose/cAMP pathway have been identified in a selection for mutants that fail to glucose repress transcription of the fbp1+ gene, encoding the gluconeogenic enzyme fructose-1,6-bisphosphatase (Hoffman and Winston 1990). This collection of mutants includes 31 strains carrying mutations in the git2+/cyr1+ adenylate cyclase gene, some of which compose two intragenic complementation groups (Hoffman and Winston 1990, 1991). In vitro assays of adenylate cyclase activity showed wild-type catalytic activity in git2-7 and git2-210 strains from one group and reduced catalytic activity in a git2-61 strain from the second group (Hoffman and Winston 1991). Presumably, the first group is defective in a step involved in in vivo activation, such as binding an activator, while the second group is defective in catalytic activity.

In this study, we carried out a suppressor selection on two activation-defective adenylate cyclase mutant strains in an effort to identify gain-of-function mutations in a gene presumed to encode an adenylate cyclase activator. From 120 independently isolated mutants from two parental strains, we identified two dominant mutations that failed to suppress an adenylate cyclase deletion. Furthermore, a synthetic interaction with a gain-of-function gpa2R176H allele suggested that these mutations affected a gene that acted in concert with Gpa2 to activate adenylate cyclase. Screening of a genomic library, constructed with insert DNA from a strain carrying one of these dominant mutations, failed to identify the suppressor gene. Surprisingly, a genetic mapping approach revealed that these suppressors are alleles of the cgs2+/pde1+ cAMP phosphodiesterase gene (DeVoti et al. 1991; Mochizuki and Yamamoto 1992). Similar to the alleles identified in this study, the cgs2-2 allele, which we show to possess a frameshift mutation, also behaves as a dominant suppressor of mutations in git genes, although the cgs2-2 allele is able to suppress a gpa2 disruption. Therefore, these appear to be dominant-negative mutations with dominance possibly due to protein poisoning or haplo-insufficiency. Measurement of cAMP levels in a cgs2-s1 strain shows that these cells possess wild-type basal cAMP levels, but exhibit a dramatically increased glucose-stimulated response indicative of a loss of feedback regulation. These data are consistent with a model in which the wild-type Cgs2 cAMP phosphodiesterase becomes activated almost immediately after adenylate cyclase activation to limit the cAMP response to glucose in fission yeast.

MATERIALS AND METHODS

Yeast strains and growth media:

Yeast strains used are listed in Table 1. The fbp1∷ura4+ and ura4∷fbp1-lacZ reporter constructs have been previously described (Hoffman and Winston 1990). Only relevant genotypes are given in the text, tables, and figure legends.

TABLE 1.

Strain list

Strain Genotype
CHP7 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-7
CHP14 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14
CHP216 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-216
CHP384 h+ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ cgs2-2
CHP386 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-2∷his7+
CHP398 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-2∷his7+
CHP732 h−ade6-M216 leu1-32 ura4∷fbp1-lacZ cgs2-s1 gpa2R176H
CHP733 h−ade6-M216 leu1-32 ura4∷fbp1-lacZ cgs2-s1
CHP769 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+cgs2-s1 git3Δ∷kan
CHP774 h+ade6-M216 his7-366 leu1-32 ura4-D18 fbp1∷ura4+git1-1 cgs2-s1
CHP863 h−ade6-M216 his3-D1 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+gpa2∷his3+
CHP934 h−his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3Δ∷kan
CHP939 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ git3Δ∷kan
FWP17 mat2-102 lys1-131 ura4-294
FWP72 h−leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+
FWP101 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+
FWP110 h+ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git1-1
FWP111 h+ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git1-1
FWP112 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+
FWP114 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-7
FWP188 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-1∷LEU2+
FWP190 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-1∷LEU2+
KGP5 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-7 cgs2-s1
KGP6 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-216 cgs2-s4
KGP7 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-1∷LEU2+cgs2-s1
KGP8 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-1∷LEU2+cgs2-s4
KGP9 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2+cgs2-s4
KGP10 h+ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2+cgs2-s1
KGP11 h+ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2-7 gpa2-249
LWP20 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14c1676-890∷LEU2+
LWP28 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14 cgs2-s1c1676-890∷LEU2+
LWP30 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14 cdc11-123
LWP31 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14 cgs2-s1 snf1Δ∷kan
LWP38 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14 cgs2-s1c1676-804∷LEU2+
LWP39 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14 cgs2-s1c1676-751∷LEU2+
LWP40 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14 cgs2+int∷LEU2+
LWP41 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3-14 cgs2-s1 cgs2+int∷LEU2+
LWP96 h+ade6-M216 his3-D1 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+cgs2-2
LWP98 h+ade6-M216 his3-D1 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+cgs2-2 gpa2∷his3+
LWP99 h+ade6-M216 his3-D1 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+
LWP110 h−leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git2+-TAP∷kan
LWP127 mat2-102 ade6-M210 ura4∷fbp1-lacZ fbp1∷ura4+git2+-13myc∷kan
LWP156 mat2-102 his3-D1 ura4∷fbp1-lacZ fbp1∷ura4+git3Δ∷kan
LWP159 h−ade6-M216 his7-366 leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+cgs2-2 git3Δ∷kan
LWP161 h+ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ cgs2-2 git3Δ∷kan
LWP167 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ cgs2-s1 git3Δ∷kan
LWP181 h+ade6-M216 leu1-32 ura4∷fbp1-lacZ cgs2+int∷LEU2+gpa2∷ura4+
LWP191 h+ade6-M216 leu1-32 ura4∷fbp1-lacZ cgs2-s1 gpa2∷ura4+
LWP238 h−ade6-M210 his7-366 leu1-32 ura4∷fbp1-lacZ cgs2-s4
RWP1 h−ade6-M216 leu1-32 ura4∷fbp1-lacZ gpa2R176H
RWP9 h−leu1-32 ura4∷fbp1-lacZ fbp1∷ura4+git3Δ∷kan
SP578 h90ade6-M216 leu1-32 cgs2-2
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Standard rich media yeast extract agar (YEA) and YEL (Gutz et al. 1974) were supplemented with 2% casamino acids. S. pombe minimal media (PM; Watanabe et al. 1988) were supplemented with required nutrients at 75 mg/liter, except for leucine, which was added to 150 mg/liter. Glucose was generally present at a concentration of 3%, unless otherwise specified. Sensitivity to 5-fluoroorotic acid (5-FOA) was determined on SC solid media containing 8% glucose as previously described (Hoffman and Winston 1991). Strains were grown at 30°.

Recombinant DNA methodology:

Standard recombinant DNA techniques, including DNA restriction digests, ligations, and Escherichia coli transformations, were done according to Ausubel et al. (1998).

Isolation and genetic analysis of git2-7 and git2-216 suppressor mutations:

5-FOA-resistant (5-FOAR) derivatives from strains CHP7 (git2-7) and CHP216 (git2-216) were isolated by plating 105 cells onto YEA plates, subjecting them to 175 J of UV irradiation, and replica plating to 5-FOA medium after a 24-hr grow out period. Colonies that formed within 3–6 days were single colony purified on YEA and retested for 5-FOAR growth. Sixty 5-FOAR derivatives from each strain were isolated and characterized. Strains unable to grow on PM-ura medium were discarded as these most likely carry mutations in either ura4+ (within the fbp1-ura4+ reporter) or ura5+ and are not strains with reduced fbp1-ura4+ transcription. (PM-ura medium is less restrictive than SC-ura for growth of strains carrying a functional fbp1-ura4+ reporter.) Random spore analysis was carried out on progeny from CHP7 derivatives crossed with CHP386 (git2Δ∷his7+) or from CHP216 derivatives crossed with CHP398 (git2Δ∷his7+). Intragenic suppressor mutations were identified by the fact that all His− progeny were 5-FOAR, while all His+ progeny were 5-FOA sensitive (5-FOAS). Suppressor mutations able to suppress git2Δ∷his7+ were identified by the fact that approximately half of the His+ (git2Δ) progeny were 5-FOAR. Allele-specific suppressors were identified by the fact that approximately half of the His− progeny, containing the git2 point mutations, were 5-FOAS, while all of the His+ progeny were 5-FOAS. The allele-specific nature of the suppressor mutations in strains KGP5 and KGP6 was confirmed by tetrad dissection. Dominance-recessiveness testing was carried out by mating the suppressor strains with a git2Δ strain and determining whether the resulting diploid strain was 5-FOAS (indicating a recessive suppressor) or 5-FOAR (indicating a dominant suppressor).

Construction and screening of an S. pombe genomic DNA library:

Genomic DNA from strain KGP5 (git2-7 sog1-1) was isolated and used as insert DNA for the construction of a genomic library in the his7+ cloning vector pEA500 (Apolinario et al. 1993), using a partial fill-in cloning strategy as described by Ohi et al. (1996). The insert DNA was generated by a Sau3AI partial digestion and partial Klenow fill-in, while the plasmid was linearized at the XhoI site in the polylinker followed by a partial Klenow fill-in reaction. The library consists of ∼250,000 independent clones, of which 70% contain inserts with an average size of 3.9 kb. CHP7 (git2-7; 50,000 colonies screened) or FWP110 (git1-1; 40,000 colonies screened) transformants were screened for 5-FOAR growth, indicative of glucose repression of fbp1-ura4+ expression. Candidate transformants were subjected to a plasmid-loss experiment to determine whether or not the 5-FOAR growth was plasmid conferred. Plasmids shown to confer 5-FOAR growth were rescued into E. coli (Hoffman and Winston 1987) and subjected to a DNA sequence analysis using vector-specific sequencing primers to determine the two ends of the insert DNA.

Mapping of sog1-1 to the cgs2+ locus:

Chromosomal mapping of sog1-1 to chromosome 3 was carried out by benomyl-induced haploidization of an h−/mat2-102 diploid strain as previously described (Alfa et al. 1993). The sog1-1 allele was further mapped by tetrad dissection, scoring for its ability to suppress either a git3-14 or a git3Δ allele. Initial crosses involved strains carrying mutant alleles of chromosome 3 genes, including ura4+, wee1+, cdc21+, ade6+, git3+, cdc11+, and ade5+. Additional markers were constructed by cloning fragments (∼1 kb) of chromosome 3 into the LEU2+-marked TOPO cloning vector pNMT41 (Invitrogen, Carlsbad, CA), linearizing the plasmid within the insert DNA, and integrating the plasmid into the chromosome by homologous recombination (Bähler et al. 1998) to place the LEU2+ marker at specific sites within the chromosome.

Gap repair cloning of the cgs2 allele from wild-type and mutant strains:

The cgs2 alleles from various strains were cloned by gap repair to avoid PCR-based amplification artifacts. Two sets of PCR primers were used to amplify regions flanking the cgs2 gene and were joined into a single product in a two-step PCR reaction (Pearson et al. 1998). Primers cgs2gap1 (5′-TTATTTGGAAATGGAGAGTCACG-3′) and cgs2gap2 (5′-ACGGGATCCAGGAGCCTTTGCTGCAGCGTG-3′) amplify a 330-bp fragment ∼800 bp downstream of the cgs2 STOP codon, while primers cgs2gap3 (5′-CCTGGATCCCGTTAGCAGGCATGGAGTGCA-3′) and cgs2gap4 (5′-AAACATCAAGAAATGGAGACTCG-3′) amplify a 260-bp fragment ∼1.7 kb upstream of the cgs2 START codon. Primers cgs2gap2 and cgs2gap3 contain complementary sequences (underlined in the sequences shown above) that include a unique BamHI site, allowing the two PCR products to be combined into a single product by a second PCR reaction. This second-round PCR product was cloned into the pNMT41 TOPO cloning vector to create a plasmid that can be linearized with BamHI to facilitate gap repair cloning of cgs2 alleles from the yeast chromosome. The cgs2 allele was cloned from strains FWP112 (cgs2+), KGP10 (sog1-1/cgs2-s1), KGP9 (sog1-4/cgs2-s4), and SP578 (cgs2-2) and the plasmids were rescued into E. coli by the “smash and grab” method (Hoffman and Winston 1987). The sequence of each cgs2 allele was determined on both strands using custom oligonucleotides together with the CEQ DTCS-Quick Start kit (Beckman Coulter) and the CEQ 8000 genetic analysis system.

β-Galactosidase assays:

Strains were cultured overnight under repressing conditions (8% glucose) in YEL medium before subculturing into YEL medium under repressing or derepressing conditions (0.1% glucose plus 3% glycerol). Cultures were grown for 24 hr to a final cell density of ∼1 × 107 cells/ml. Protein lysates were prepared on ice and assayed for β-galactosidase activity as previously described (Nocero et al. 1994).

cAMP assays:

Intracellular cAMP levels were measured by radioimmune assay in glucose-starved cells (basal level) and in the same cultures 1 and 10 min after exposure to 100 mm glucose as previously described (Byrne and Hoffman 1993).

RESULTS

Isolation of allele-specific adenylate cyclase suppressor mutations:

In an effort to identify direct activators of S. pombe adenylate cyclase, the following suppressor screen was carried out. Strains CHP7 (git2-7) and CHP216 (git2-216) carry mutations in the adenylate cyclase gene that display intragenic complementation with the git2-61 mutation, which confers a partial defect in catalytic activity (Hoffman and Winston 1991). As such, the git2-7 and git2-216 mutations may affect the binding of an activator. These mutations confer a 5-FOAS growth defect (Figure 1; the 5-FOAS phenotype conferred by the git2-216 mutation is leaky, although sufficient to allow selection of suppressor mutations) due to derepression of transcription of an fbp1-ura4+ reporter gene, which normally is glucose repressed. Sixty independently isolated 5-FOAR derivatives from each of strains CHP7 and CHP216 were isolated and put through genetic tests to identify candidate gain-of-function mutations in an adenylate cyclase activator gene (see materials and methods). One suppressor mutation from each screen displayed the desired genetic characteristics of being unlinked to the original git2 mutation, being dominant to wild type for suppression of the git2 point mutations, and being unable to suppress a git2 deletion (git2Δ; Figure 1). These mutations, designated sog for suppressor of git, are single nuclear mutations, segregating 2:2 in 18 tetrads each. Finally, these mutations can suppress either the git2-7 or the git2-216 mutation and are tightly linked to each other, suggesting that they are allelic, as a cross between strains KGP5 (git2-7 sog1-1) and KGP6 (git2-216 sog1-4) yielded only 5-FOAR progeny in 38 tetrads.

Figure 1.

Figure 1.

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sog1-1 and sog1-4 mutations suppress spontaneous mutations, but not deletions, of the git2+/cyr1+ adenylate cyclase gene. Strains were spotted onto YEA-rich medium and grown overnight before replica plating to YEA, 5-FOA, and SC-Ura media. Plates were photographed after 2 days. Glucose repression of the fbp1-ura4+ reporter confers 5-FOAR growth, while a defect in glucose repression allows strains to grow on SC-Ura as previously shown (Hoffman and Winston 1990). Note that the git2-216 allele allows some growth on 5-FOA medium, although this amount of growth did not preclude our ability to carry out a screen for suppressor mutations that allows single-colony formation on 5-FOA. Suppression of the git1-1 mutation by sog1-1 is also shown.

Genetic interactions between sog1-1 and “upstream” git genes:

To determine if sog1-1 is a gain-of-function allele of any of the known git genes that are required for adenylate cyclase activation, we crossed strain KGP10 (git2+ sog1-1) with strains carrying a mutation in git1+, git3+, git5+, git7+, gpa2+, git10+, or git11+. With the exception of the cross with a gpa2− mutant strain, most tetrads from each cross contained three 5-FOAR to one 5-FOAS progeny, indicating that the sog1-1 allele is not linked to the git mutation in the other parental strain and that the sog1-1 allele suppresses the git mutant allele in the double-mutant progeny (see Figure 1 for suppression of git1-1 by sog1-1). On the other hand, every tetrad from the gpa2− mutant cross contained two 5-FOAR progeny and two 5-FOAS progeny, indicating that sog1-1 either is tightly linked to the gpa2 locus or fails to suppress a gpa2− mutation. To distinguish between these two possibilities, progeny from a cross between KGP11 (git2-7 gpa2-249) and KGP5 (git2-7 sog1-1) were analyzed. Tetrads with fewer than two 5-FOAR progeny were readily obtained (data not shown), indicating that git2-7 gpa2-249 sog1-1 5-FOAS progeny were being generated. Thus, sog1-1 fails to suppress a mutation in the gpa2+ gene that encodes the Gα of the heterotrimeric G protein of the glucose/cAMP pathway (Nocero et al. 1994), but does suppress mutations in the other git genes required for adenylate cyclase activation. In addition, strains carrying the sog1-1 allele together with the activated gpa2R176H allele display a synthetic defect in fbp1-lacZ derepression (see below), suggesting that Sog1 and Gpa2 may act in concert to activate adenylate cyclase.

Efforts to clone sog1-1 from a genomic library identify only multicopy suppressors:

As the sog1-1 mutation is dominant (i.e., a git2-7/git2Δ sog1-1/sog1+ diploid is 5-FOAR), we constructed a genomic library from strain KGP5 (git2-7 sog1-1) and screened it for plasmids able to confer 5-FOAR growth to either CHP7 (git2-7) or CHP110 (git1-1) transformants (see materials and methods). Surprisingly, all nine candidate plasmids that suppress the git1-1 mutation carry the pka1+ gene, which encodes the catalytic subunit of the cAMP-dependent protein kinase (Table 2). The plasmids obtained from the git2-7 transformants carried a more varied collection of genes, including pka1+, sck2+, pyp1+, mcs4+, spc1+, and esc1+. However, it seemed unlikely that these clones carried sog1-1, as they also confer 5-FOAR growth to strains carrying deletions of either git2+ or pka1+ (see Figure 2 for suppression by esc1+). In addition, homologous integration of the plasmids, followed by a linkage analysis, demonstrated that none of these genes are linked to sog1+; therefore these genes represent multicopy suppressors.

TABLE 2.

Multicopy suppressors identified from sog1-1 library

No. of clones
Gene cloned Suppressor of git1-1 Suppressor of git2-7
pka1+ 9 1
esc1+ 0 4
pyp1+ 0 1
mcs4+ 0 3
sck2+ 0 2
spc1+/sty1+ 0 1
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Figure 2.

Figure 2.

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Multicopy suppression of git2− and pka1− mutations by overexpression of esc1+. His+ transformants carrying either the pEA500 empty vector (Apolinario et al. 1993) or pYZ3 (esc1+) were pregrown on PM-his medium before replica plating to PM-his and to 5-FOA media. Plates were photographed after 3 days.

sog1-1 maps to the cgs2+ locus:

Due to the inability to clone sog1-1 by a library screen, we adopted a genetic mapping strategy to identify the sog1+ gene. We first mapped sog1-1 to chromosome 3 using a diploid haploidization strategy (see materials and methods), which takes advantage of the mat2-102 mating-type allele to construct stable diploids by mating a mat2-102 strain with an h− strain. Benomyl treatment of diploid cells formed by mating strains FWP17 and KGP5, which carry markers to identify all three chromosomes from each parental strain, resulted in haploid derivatives without undergoing meiotic recombination so that markers on each chromosome remain genetically linked. Of the haploid strains that carried chromosome 2 from the KGP5 parent (with the git2-7 and fbp1-ura4+ alleles), only the strains carrying chromosome 3 from the sog1-1 parental strain were 5-FOAR, due to the suppression of the git2-7 mutation by sog1-1 (data not shown). Thus, the sog1+ gene is present on chromosome 3.

A series of tetrad dissections involving strains with various chromosome 3 markers led us to the cgs2+ cAMP phosphodiesterase gene (Figure 3; additional crosses involving markers that are unlinked to the sog1-1 mutant allele are not shown). These mapping experiments involved the use of available chromosome 3 markers, along with engineered markers created by homologous integration of a LEU2-marked plasmid at various sites along the chromosome (see materials and methods). One such marker, integrated to within 1.5 kb of the cgs2+ translational start site, maps to within 0.45 cM of sog1-1. Thus, it appears that sog1-1 is either an allele of cgs2+ or a gene adjacent to cgs2+.

Figure 3.

Figure 3.

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Genetic mapping of sog1-1 on chromosome 3. (A) Schematic of a portion of chromosome 3 displaying the relative locations of the sog1-1 allele, the snf1+/ssp2+ gene (systematic name SPCC74.03c), the cdc11+ gene, and four LEU2+-marked plasmid integrations (see materials and methods). The cgs2int integration site is within 1.5 kb of the translational start of the cgs2+ gene, while the L751, L804, and L890 integration sites are at approximately positions 751,000, 804,000, and 890,000, respectively, of contig c1676. Distances from sog1-1 are given both in kilobase pairs and in centimorgans (as determined from our genetic mapping experiments). (B) Linkage data used to determine genetic map distances from sog1-1.

sog1-1 and sog1-4 are alleles of the cgs2+ cAMP phosphodiesterase gene:

To determine whether sog1-1 and sog1-4 are alleles of cgs2, we cloned and sequenced cgs2 from sog1-1 and sog1-4 strains, as well as from cgs2+ and cgs2-2 strains. These alleles were cloned by gap repair to avoid PCR amplification errors (see materials and methods ). DNA sequence analyses of the cloned alleles showed that the cgs2-2 mutation is a +1 frameshift in codon 270 (a duplication of the T at nucleotide 1366 in the genomic sequence; the cgs2+ gene contains three introns with a total length of 559 nucleotides), leading to a truncated product of 273 residues. The cgs2 allele from a sog1-1 strain contains a single-point mutation (C to T) at nucleotide 71, changing residue 24 from serine to phenylalanine (Figure 4A). The cgs2 allele from a sog1-4 strain has a single-point mutation (G to A) at nucleotide 1365, changing residue 269 from glycine to aspartic acid (Figure 4B). These sequence changes were confirmed by PCR amplification and direct sequencing of the PCR products from strains KGP5 (git2-7 sog1-1) and KGP6 (git2-216 sog1-4). As shown in Figure 4, the residues affected by these mutations are found within two highly conserved sequences present in phosphodiesterases from a wide variety of organisms, including other yeasts and fungi (S. cerevisiae, Kluyveromyces lactis, Debaryomyces hansenii), human and plant fungal pathogens (Candida albicans, C. glabrata, Cryptococcus neoformans, Ashbya gossypii), and bacterial pathogens (Yersinia pestis, Legionella pneumophila, Vibrio fischeri). In a complete alignment of these 11 enzymes, only 22 residues are perfectly conserved (data not shown). Thus the presence of 12 of these conserved residues in these two small regions of the proteins suggests that we have identified two conserved domains that are important for phosphodiesterase function and/or regulation and that the sog1-1 and sog1-4 mutations reduce phosphodiesterase activity to suppress defects in the glucose/cAMP pathway. Thus sog1-1 and sog1-4 are two mutant alleles of cgs2 and have been renamed cgs2-s1 and cgs2-s4.

Figure 4.

Figure 4.

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cgs2-s1 and cgs2-s4 mutations alter residues in highly conserved domains within the Cgs2 protein. (A) Alignment of N-terminal domains that include the residue affected by the cgs2-s1 mutation from phosphodiesterases from S. pombe (NP_588337), S. cerevisiae (NP_011266), K. lactis (XP_453575), D. hansenii (XP_460986), C. albicans (P32782), C. glabrata (XP_447131), C. neoformans (XP_568076), A. gossypii (NP_985806), Y. pestis (NP_406214), L. pneumophila (YP_095961), and V. fischeri (Q56686). The number preceding the protein sequence indicates the position of the residue that aligns with serine 24 of Cgs2, which is changed to phenylalanine (this position is shown above the alignment) in the Cgs2-s1 protein. (B) Alignment of the conserved domain containing glycine 269 of Cgs2 with the same phosphodiesterases as in A. The number preceding the protein sequence indicates the position of the residue that aligns with glycine 269 of Cgs2, which is changed to aspartic acid (this position is shown above the alignment) in the Cgs2-s4 protein. Note that the cgs2-2 mutation is a frameshift in codon 270.

Comparison of cgs2-s1 and cgs2-2 mutations:

The demonstration that the sog1-1 and sog1-4 suppressor mutations are alleles of the cgs2+ cAMP phosphodiesterase gene was unexpected for two reasons. We expected to obtain cgs2 loss-of-function alleles among our suppressors of the adenylate cyclase defect, as the loss of phosphodiesterase activity would elevate cAMP levels; however, we assumed that such alleles would be recessive to wild type. In addition, we assumed that such cgs2− mutations would be able to suppress gpa2− Gα mutations, as gpa2− mutants display a measurable basal cAMP level (Isshiki et al. 1992; Byrne and Hoffman 1993). We therefore revisited these observations by comparing cgs2-s1 with the previously identified cgs2-2 allele both for dominance and for suppression of gpa2− mutations. Both cgs2-s1 and cgs2-2 behave as dominant suppressors of a git3 deletion (Welton and Hoffman 2000) as judged by the 5-FOAR growth of diploid strains that are homozygous git3Δ and heterozygous at the cgs2 locus (Figure 5). In fact, the cgs2-2/cgs2+ diploid appears to be more resistant to 5-FOA than the cgs2-s1/cgs2+ diploid. A quantitative assessment of suppression by measuring β-galactosidase activity expressed from the fbp1-lacZ reporter confirms this observation (Table 3). In haploid git3Δ strains, the cgs2-2 allele completely restores glucose-repressed levels under repressing conditions and confers a significant defect in derepression under glucose-starved conditions. The cgs2-s1 allele significantly reduces fbp1-lacZ expression under repressing conditions, but has only a modest effect on derepressed levels of expression. Diploid cgs2-2/cgs2+ cells express the fbp1-lacZ reporter at the same levels as the cgs2-2 haploid strain does; thus the cgs2-2 allele is dominant to the wild-type allele with respect to the regulation of fbp1-lacZ and fbp1-ura4 transcription. Similar to its ability to confer 5-FOAR growth (Figure 5), the cgs2-s1 allele restores repression of fbp1-lacZ expression even in the presence of the wild-type cgs2+ allele, although not to the same extent as the cgs2-2 allele (Table 3).

Figure 5.

Figure 5.

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cgs2-s1 and cgs2-2 mutations are dominant suppressors of a deletion of the git3+ GPCR gene. Haploid and diploid strains were pregrown on YEA medium overnight at 30° before replica plating to YEA and 5-FOA media. Plates were photographed after 3 days.

TABLE 3.

Suppression of git3Δ by cgs2-s1 and cgs2-2 in haploid and diploid strains

β-Galactosidase activity
Strain Genotype Repressed Derepressed
FWP72 git3+cgs2+ 3 ± 0 1369 ± 142
RWP9 git3Δ cgs2+ 925 ± 10 1282 ± 10
LWP167 git3Δ cgs2-s1 27 ± 10 527 ± 56
LWP161 git3Δ cgs2-2 4 ± 0 52 ± 20
LWP127 × LWP110 git3+/git3+cgs2+/cgs2+ 6 ± 1 1754 ± 32
LWP156 × CHP934 git3Δ/git3Δ cgs2+/cgs2+ 513 ± 130 1573 ± 32
LWP156 × CHP769 git3Δ/git3Δ cgs2+/cgs2-s1 68 ± 15 970 ± 234
LWP156 × LWP159 git3Δ/git3Δ cgs2+/cgs2-2 4 ± 1 43 ± 22
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β-Galactosidase activity was determined from three independent cultures as described in materials and methods. The average ±SE represents specific activity per milligram of soluble protein.

We next examined the ability of cgs2-2 and cgs2-s1 alleles to suppress mutations in the gpa2+ gene that encodes a Gα subunit, which we have shown to directly bind to an N-terminal domain of adenylate cyclase (Ivey and Hoffman 2005). Using two different marked disruptions of gpa2+ that result in similar levels of fbp1-lacZ expression in glucose-grown cells (Table 4), we show that cgs2-2 completely restores glucose-repressed levels of expression while cgs2-s1 has only a modest effect on expression (Table 4). These results are similar to observations using spontaneous gpa2− mutant alleles with regard to the 5-FOA growth phenotype, which reflects transcription of the fbp1-ura4 reporter. We observe that the cgs2-2 allele restores 5-FOAR growth while the cgs2-s1 allele does not (data not shown).

TABLE 4.

Differential suppression of a gpa2 deletion by cgs2-s1 and cgs2-2

Strain Genotype β-Galactosidase repressed
LWP99 gpa2+cgs2+ 7 ± 1
CHP863 gpa2∷his3+cgs2+ 1351 ± 48
LWP181 gpa2∷ura4+cgs2+ 1221 ± 55
CHP733 gpa2+cgs2-s1 4 ± 0
LWP96 gpa2+cgs2-2 7 ± 1
LWP191 gpa2∷ura4+cgs2-s1 482 ± 71
LWP98 gpa2∷his3+cgs2-2 10 ± 1
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β-Galactosidase activity was determined from three independent cultures as described in materials and methods. The average ±SE represents specific activity per milligram of soluble protein. The gpa2∷ura4+ disruption allele was previously described (Isshiki et al. 1992), while the gpa2∷his3+ allele was created by the nonhomologous integration of plasmid pAF1 into codon 329 of the 354-codon open reading frame (Hoffman and Welton 2000).

To investigate the roles of Cgs2 and Gpa2 in the glucose/cAMP pathway further, we examined fbp1-lacZ expression in double-mutant strains carrying either cgs2-2 or cgs2-s1 in combination with gpa2R176H, an allele activated due to a loss of GTPase activity. We had previously shown that gpa2R176H suppresses the loss of the Git3 GPCR, the Git5 Gβ subunit, or the Git11 Gγ subunit, indicating that the role of the Gβγ dimer is to couple the Gpa2 Gα to the Git3 GPCR, so that, upon glucose detection, Git3 could promote the activation of Gpa2 (Welton and Hoffman 2000). We had also noted that the gpa2R176H allele confers only a partial defect in fbp1-lacZ derepression. The cgs2-s1 or gpa2R176H single-mutant strains display a moderate reduction in fbp1-lacZ derepression, while the cgs2-s1 gpa2R176H double mutant displays an almost total loss of derepression (Table 5). On the other hand, the cgs2-2 allele is sufficient to confer a severe defect in fbp1-lacZ derepression. The synthetic interaction between the cgs2-s1 or gpa2R176H alleles supports a model in which cAMP phosphodiesterase activity serves as a counterbalance to Gpa2-mediated adenylate cyclase activation.

TABLE 5.

Synthetic defect in fbp1-lacZ derepression by cgs2-s1 and gpa2R176H

β-Galactosidase activity
Strain Genotype Repressed Derepressed
FWP72 gpa2+cgs2+ 3 ± 0 1369 ± 142
CHP733 gpa2+cgs2-s1 4 ± 0 263 ± 12
RWP1 gpa2R176Hcgs2+ 2 ± 0 483 ± 5
CHP732 gpa2R176Hcgs2-s1 3 ± 0 12 ± 3
CHP384 gpa2+cgs2-2 3 ± 0 24 ± 2
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Cells were grown in PM complete medium and β-galactosidase activity was determined from three independent cultures as described in materials and methods. The average ±SE represents specific activity per milligram of soluble protein.

cgs2-s1 and cgs2-s4 strains display distinct cAMP responses to glucose:

To examine the effect of the cgs2-s1 and cgs2-s4 mutations on cAMP signaling, we assayed the glucose-triggered cAMP response in wild-type cells and in various mutant strains (Figure 6). Similar to our previous observations (Byrne and Hoffman 1993), wild-type S. pombe cells respond to glucose exposure with a four- to sixfold increase in cAMP levels, while cgs2-2 cells display an elevated basal cAMP level and a similar-fold increase in cAMP levels after glucose addition (Figure 6). The cgs2-s4 cells display an intermediate result with cAMP levels that are elevated relative to wild-type cells, but not to the same degree as cgs2-2 cells are, at each time point. Overall, the cgs2-s4 response resembles that of the cgs2-2 cells in that cAMP levels accumulate at a somewhat linear rate over the course of the 10-min assay (this includes data from a 5-min time point; data not shown). Remarkably, the cgs2-s1 cells possess a basal cAMP level similar to that of wild-type cells; however, within 1 min of glucose exposure, the cAMP levels reach that observed in the glucose-stimulated cgs2-2 cells. More importantly, whereas the rate of cAMP accumulation in wild-type cells decreases significantly after the 1-min time point, the rate of cAMP accumulation in cgs2-s1 cells is only moderately reduced, leading to a total accumulation at the 10-min time point exceeding that of the cgs2-2 cells. However, the cgs2-s1 response profile does resemble that of the cgs2+ cells in that it is biphasic with a rapid increase in cAMP levels in the first minute, followed by a reduced rate of cAMP accumulation during the remaining 9 min. The significance of the linear vs. the biphasic response curves is discussed below.

Figure 6.

Figure 6.

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cAMP response curves in wild-type and mutant strains. Cells were cultured for 24 hr in medium containing 0.1% glucose and 3% glycerol. cAMP levels were measured as previously described (Byrne and Hoffman 1993) immediately prior to glucose addition to the cultures, as well as 1 min and 10 min after glucose addition. Strains carried either the wild-type git3+ gene (solid lines) or a git3Δ deletion allele (dashed lines), as well as the wild-type cgs2+ gene (circles), the cgs2-s1 allele (triangles), the cgs2-s4 allele (diamonds), or the cgs2-2 allele (squares).

We carried out parallel cAMP assays on git3Δ strains to better understand how the cgs2-s1 mutation suppresses the loss of the Git3 receptor protein. The git3Δ cgs2+ cells display both a reduced basal cAMP level and a loss of the glucose-triggered cAMP response (Figure 6). The git3Δ cgs2-s1 cells possess a basal cAMP level similar to that of wild-type cells, but show little or no response to glucose. The git3Δ cgs2-2 cells possess significantly elevated basal cAMP levels that are largely unaffected by glucose addition. These assays demonstrate that the cgs2-s1 mutation does not suppress the git3Δ allele by restoring adenylate cyclase activation, but does increase basal cAMP levels enough to restore glucose repression of fbp1+ transcription.

DISCUSSION

The original goal of this study was to identify a direct activator of S. pombe Git2/Cyr1 adenylate cyclase enzyme by isolating allele-specific suppressors of git2-7 and git2-216 alleles that produce activation-defective enzymes. We assumed that these suppressor mutations would restore an interaction between adenylate cyclase and the activator protein and would thus be dominant. We unexpectedly found that loss-of-function alleles of the cgs2+ cAMP phosphodiesterase gene are dominant for suppression of the git2 mutant alleles (Figure 5, Table 3). This dominant-negative phenotype could be due to either haplo-insufficiency or a protein-poisoning effect of the defective protein by forming inactive multimeric complexes with wild-type subunits.

As cgs2-s1 and cgs2-s4 are dominant suppressor mutations, we tried to clone cgs2-s1 from a genomic library containing insert DNA from KGP5 (git2-7 cgs2-s1) cells. While failing to identify cgs2-s1, our screen identified several interesting multicopy suppressors (Table 2). We repeatedly cloned pka1+, recovering seven distinct clones whose inserts ranged from 3.4 to 8.4 kb, demonstrating the high quality of this his7+-marked library. The remaining multicopy suppressors encode proteins that fall into three groups. The Sck2 kinase (Fujita and Yamamoto 1998) is 42% identical to S. pombe Sck1, which we previously identified as a multicopy suppressor of PKA pathway mutations (Jin et al. 1995). Mcs4, Pyp1, and Spc1 are components of a stress-activated MAP kinase (SAPK) pathway, required for fbp1+ transcription. The Pyp1 protein tyrosine phosphatase dephosphorylates and inactivates the Spc1 SAPK (Millar et al. 1995; Shiozaki and Russell 1995). We previously showed that pyp1+ and the closely related pyp2+ gene are multicopy suppressors of PKA pathway mutants (Dal Santo et al. 1996). It is less obvious why Spc1 or the Mcs4 response regulator were identified in this screen. Spc1 overexpression may create a pool of inactive protein that inhibits activation of a target protein required for fbp1+ transcription such as the Atf1 bZIP transcriptional activator (Shiozaki and Russell 1996; Gaits et al. 1998). Similarly, Mcs4 overexpression may produce partially assembled signaling complexes that reduce Spc1 activation. This is not the first instance in which positive and negative regulators of a S. pombe MAP kinase (MAPK) pathway have been found to confer the same multicopy suppressor phenotype. The Pek1 MAPK kinase and the Pmp1 MAPK phosphatase, which antagonistically regulate the Pmk1 cell wall integrity MAPK, are multicopy suppressors of the chloride-sensitive growth conferred by disruption of the ppb1+ calcineurin gene (Sugiura et al. 1998, 1999). As signaling from both MAPK pathways is reduced by overexpression of positive regulators, the proper stoichiometry of pathway components appears to be important for signaling. Finally, the Esc1 helix-turn helix protein was previously identified as a multicopy suppressor of a temperature-sensitive lethal mutation in pat1+ (pat1-114), which encodes a protein kinase that regulates meiotic entry (Benton et al. 1993). Interestingly, increased PKA activity due to loss of the Cgs1 PKA regulatory subunit or the Cgs2 PDE also suppresses pat1-114 (DeVoti et al. 1991), as does loss of SAPK activation (Stettler et al. 1996). Therefore, overexpression of Esc1 may mimic either a “high-PKA-activity” or a “low-SAPK-activity” phenotype.

Prior to this study, we assumed that the cAMP response to glucose was a function of adenylate cyclase regulation while Cgs2 PDE activity remained constant. This was due to the observation that cgs2-2 cells display a similar-fold increase in cAMP levels as seen in wild-type cells, although the increase in absolute cAMP levels is significantly higher in cgs2-2 cells (Byrne and Hoffman 1993). However, the cAMP response data in Figure 6 challenge this model. The rate of cAMP accumulation in both cgs2-2 and cgs2-s4 cells is relatively constant over the course of the 10-min assay (this is further supported by 5-min time points taken for the cgs2-s4 cells; data not shown). In contrast, the rate of cAMP accumulation drops off significantly after the first minute in both cgs2+ and cgs2-s1 cells. These results are consistent with our previous assays of cgs2+ and cgs2-2 strains, which were carried out over a 2-hr time course. In these assays, cAMP accumulation in cgs2-2 cells is constant for the first 20 min after glucose addition, while the rate of cAMP accumulation in cgs2+ cells falls 10-fold after the first minute (Byrne and Hoffman 1993). The cgs2-2 mutation is a frameshift that removes the carboxy-terminal 20% of Cgs2, most likely inactivating the enzyme. The resulting linear response to glucose seen in Figure 6, as compared with the biphasic response seen for the wild-type cells, suggests that in wild-type cells, PDE becomes activated shortly after adenylate cyclase. The linear response and intermediate cAMP values observed in cgs2-s4 cells is consistent with a severe reduction, but not a total loss, of PDE activity. Meanwhile, the basal cAMP level in cgs2-s1 cells is similar to that of wild-type cells, while the glucose-stimulated level resembles that of cgs2-2 cells. The serine-to-phenylalanine substitution at residue 24 in the Cgs2-s1 protein occurs in one of the most highly conserved regions of this family of phosphodiesterases (Figure 4A). This domain may be required for PDE activation by binding cyclic nucleotides to allosterically regulate PDE activity or through a post-translational modification, similar to the S. cerevisiae Pde1 enzyme, which appears to be activated by PKA phosphorylation of serine 252 (Ma et al. 1999).

Alternatively, our original model, in which only adenylate cyclase regulation controls the cAMP response, could be correct if activation and feedback mechanisms independently regulate adenylate cyclase activity. In wild-type cells, glucose signaling through the Git3 GPCR would lead to Gpa2-mediated activation of adenylate cyclase (Welton and Hoffman 2000; Ivey and Hoffman 2005), while increased cAMP levels would trigger a feedback response to downregulate adenylate cyclase activity. In cgs2-2 cells, high basal cAMP levels would trigger the feedback mechanism without preventing glucose-mediated activation, allowing adenylate cyclase to be activated by glucose, but to a lesser extent than in wild-type cells. This would produce a linear increase in cAMP levels, as the feedback regulatory event had occurred prior to glucose detection. In this model, the Csg2-s1 PDE activity is high enough to control basal cAMP levels in glucose-starved cells, but insufficient to limit the cAMP response in glucose-stimulated cells. However, we do not favor this model since PKA activity negatively regulates the level of cAMP pathway components, as we previously showed for cgs1+ and pka1+ transcription (Stiefel et al. 2004). Both the Git3 GPCR and adenylate cyclase protein levels are significantly lower in a PKA-dependent manner in glucose-grown cells than in glucose-starved cells (D. Chandler-Militello, L. Wang and C. S. Hoffman, unpublished results). The cAMP assays are carried out after a 24-hr glucose starvation period to derepress the glucose detection apparatus. However, such derepression should not occur in cgs2-s4 and cgs2-2 cells due to their high basal cAMP levels. Therefore, the reduced rate of cAMP accumulation in these strains is most likely due to a lower concentration of adenylate cyclase at the time of glucose addition, rather than to a reduction in the degree of stimulation of the enzyme.

While characterizing genetic interactions between cgs2-s1 and mutations in other glucose/cAMP pathway genes, we uncovered a paradox with regard to the relative importance of these genes in the pathway. cgs2-s1 fails to suppress a gpa2 deletion (Table 4), but is able to suppress git1, git7, and git10 mutations, suggesting that the Gpa2 Gα subunit is more important in cAMP signaling than Git1, Git7, and Git10. However, we previously showed that the activated gpa2R176H allele cannot suppress git1, git7, or git10 mutations, suggesting that Git1, Git7, and Git10 are required for adenylate cyclase activation even when Gpa2 is mutationally activated (Welton and Hoffman 2000). In addition, git1 mutations confer a higher level of fbp1-lacZ expression in glucose-grown cells than do gpa2 mutations (Hoffman and Winston 1990, 1991; Nocero et al. 1994), suggesting that Git1 plays a more important role in cAMP signaling than Gpa2. These conflicting results may reflect differences in the roles of these genes in maintaining basal cAMP levels vs. activating adenylate cyclase or may point to an as-yet-unidentified interaction between Gpa2 and Cgs2 to regulate the production and turnover of cAMP. Such a regulatory interaction would be consistent with the observation that both gpa2R176H and cgs2-s1 strains retain some ability to regulate fbp1-lacZ expression, while the gpa2R176H cgs2-s1 double mutant is completely defective in derepression (Table 5). The Cgs2+ PDE may be able to compensate for the presumed increased adenylate cyclase activation by the Gpa2R176H subunit, while the Cgs2-s1 enzyme cannot. Further analysis of the function of the N-terminal domain of Cgs2 that is altered in the Cgs2-s1 protein may help to determine the role of this domain and whether there is a direct regulatory interaction between Cgs2 and Gpa2 to control cAMP signaling in S. pombe.

Acknowledgments

We thank Maureen McLeod, Peter Fantes, Carl Mann, and Jürg Kohli for strains carrying chromosome 3 markers. We also thank Kazahiro Shiozaki, Johan Thevelein, and Evan Kantrowitz for valuable discussions during the course of these studies. This work was supported by National Institutes of Health grant GM46226 to C.S.H.

References

  1. Alfa, C., P. Fantes, J. Hyams, M. McLeod and E. Warbrick, 1993. Experiments With Fission Yeast. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
  2. Apolinario, E., M. Nocero, M. Jin and C. S. Hoffman, 1993. Cloning and manipulation of the Schizosaccharomyces pombe his7+ gene as a new selectable marker for molecular genetic studies. Curr. Genet. 24: 491–495. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman et al., 1998. Current Protocols in Molecular Biology. Wiley Interscience, New York.
  4. Bähler, J., J. Q. Wu, M. S. Longtine, N. G. Shah, A. McKenzie, III et al., 1998. Heterologous modules for efficient and versatile PCR-based gene targeting in Schizosaccharomyces pombe. Yeast 14: 943–951. [DOI] [PubMed] [Google Scholar]
  5. Benton, B. K., M. S. Reid and H. Okayama, 1993. A Schizosaccharomyces pombe gene that promotes sexual differentiation encodes a helix-loop-helix protein with homology to MyoD. EMBO J. 12: 135–143. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Byrne, S. M., and C. S. Hoffman, 1993. Six git genes encode a glucose-induced adenylate cyclase activation pathway in the fission yeast Schizosaccharomyces pombe. J. Cell Sci. 105: 1095–1100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Dal Santo, P., B. Blanchard and C. S. Hoffman, 1996. The Schizosaccharomyces pombe pyp1 protein tyrosine phosphatase negatively regulates nutrient monitoring pathways. J. Cell Sci. 109: 1919–1925. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. DeVoti, J., G. Seydoux, D. Beach and M. McLeod, 1991. Interaction between ran1+ protein kinase and cAMP dependent protein kinase as negative regulators of fission yeast meiosis. EMBO J. 10: 3759–3768. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Fujita, M., and M. Yamamoto, 1998. S. pombe sck2+, a second homologue of S. cerevisiae SCH9 in fission yeast, encodes a putative protein kinase closely related to PKA in function. Curr. Genet. 33: 248–254. [DOI] [PubMed] [Google Scholar]
  10. Fukui, Y., T. Kozasa, Y. Kaziro, T. Takeda and M. Yamamoto, 1986. Role of a ras homolog in the life cycle of Schizosaccharomyces pombe. Cell 44: 329–336. [DOI] [PubMed] [Google Scholar]
  11. Gaits, F., G. Degols, K. Shiozaki and P. Russell, 1998. Phosphorylation and association with the transcription factor Atf1 regulate localization of Spc1/Sty1 stress-activated kinase in fission yeast. Genes Dev. 12: 1464–1473. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Gutz, H., H. Heslot, U. Leupold and N. Loprieno, 1974. Schizosaccharomyces pombe, pp. 395–446 in Handbook of Genetics, edited by R. C. King. Plenum Press, New York.
  13. Hoffman, C. S., 2005. Except in every detail: comparing and contrasting G protein signaling in Saccharomyces cerevisiae and Schizosaccharomyces pombe. Eukaryot. Cell 4: 495–503. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Hoffman, C. S., and R. Welton, 2000. Mutagenesis and gene cloning in Schizosaccharomyces pombe via nonhomologous plasmid integration and rescue. BioTechniques 28: 532–536, 538, 540. [DOI] [PubMed] [Google Scholar]
  15. Hoffman, C. S., and F. Winston, 1987. A ten-minute DNA preparation from yeast efficiently releases autonomous plasmids for transformation of Escherichia coli. Gene 57: 267–272. [DOI] [PubMed] [Google Scholar]
  16. Hoffman, C. S., and F. Winston, 1990. Isolation and characterization of mutants constitutive for expression of the fbp1 gene of Schizosaccharomyces pombe. Genetics 124: 807–816. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Hoffman, C. S., and F. Winston, 1991. Glucose repression of transcription of the Schizosaccharomyces pombe fbp1 gene occurs by a cAMP signaling pathway. Genes Dev. 5: 561–571. [DOI] [PubMed] [Google Scholar]
  18. Isshiki, T., N. Mochizuki, T. Maeda and M. Yamamoto, 1992. Characterization of a fission yeast gene, gpa2, that encodes a Gα subunit involved in the monitoring of nutrition. Genes Dev. 6: 2455–2462. [DOI] [PubMed] [Google Scholar]
  19. Ivey, F. D., and C. S. Hoffman, 2005. Direct activation of fission yeast adenylate cyclase by the Gpa2 Gα of the glucose signaling pathway. Proc. Natl. Acad. Sci. USA 102: 6108–6113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Jin, M., M. Fujita, B. M. Culley, E. Apolinario, M. Yamamoto et al., 1995. sck1, a high copy number suppressor of defects in the cAMP-dependent protein kinase pathway in fission yeast, encodes a protein homologous to the Saccharomyces cerevisiae SCH9 kinase. Genetics 140: 457–467. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Kraakman, L., K. Lemaire, P. Ma, A. W. Teunissen, M. C. Donaton et al., 1999. A Saccharomyces cerevisiae G-protein coupled receptor, Gpr1, is specifically required for glucose activation of the cAMP pathway during the transition to growth on glucose. Mol. Microbiol. 32: 1002–1012. [DOI] [PubMed] [Google Scholar]
  22. Landry, S., and C. S. Hoffman, 2001. The git5 Gβ and git11 Gγ form an atypical Gβγ dimer acting in the fission yeast glucose/cAMP pathway. Genetics 157: 1159–1168. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Landry, S., M. T. Pettit, E. Apolinario and C. S. Hoffman, 2000. The fission yeast git5 gene encodes a Gβ subunit required for glucose-triggered adenylate cyclase activation. Genetics 154: 1463–1471. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Lengeler, K. B., R. C. Davidson, C. D'Souza, T. Harashima, W. C. Shen et al., 2000. Signal transduction cascades regulating fungal development and virulence. Microbiol. Mol. Biol. Rev. 64: 746–785. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Liu, H., C. A. Styles and G. R. Fink, 1993. Elements of the yeast pheromone response pathway required for filamentous growth of diploids. Science 262: 1741–1744. [DOI] [PubMed] [Google Scholar]
  26. Lorenz, M. C., X. Pan, T. Harashima, M. E. Cardenas, Y. Xue et al., 2000. The G protein-coupled receptor Gpr1 is a nutrient sensor that regulates pseudohyphal differentiation in Saccharomyces cerevisiae. Genetics 154: 609–622. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Ma, P., S. Wera, P. Van Dijck and J. M. Thevelein, 1999. The PDE1-encoded low-affinity phosphodiesterase in the yeast Saccharomyces cerevisiae has a specific function in controlling agonist-induced cAMP signaling. Mol. Biol. Cell 10: 91–104. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Mbonyi, K., M. Beullens, K. Detremerie, L. Geerts and J. M. Thevelein, 1988. Requirement of one functional RAS gene and inability of an oncogenic ras variant to mediate the glucose-induced cyclic AMP signal in the yeast Saccharomyces cerevisiae. Mol. Cell. Biol. 8: 3051–3057. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Millar, J. B., V. Buck and M. G. Wilkinson, 1995. Pyp1 and Pyp2 PTPases dephosphorylate an osmosensing MAP kinase controlling cell size at division in fission yeast. Genes Dev. 9: 2117–2130. [DOI] [PubMed] [Google Scholar]
  30. Mochizuki, N., and M. Yamamoto, 1992. Reduction in the intracellular cAMP level triggers initiation of sexual development in fission yeast. Mol. Gen. Genet. 233: 17–24. [DOI] [PubMed] [Google Scholar]
  31. Nikawa, J., S. Cameron, T. Toda, K. M. Ferguson and M. Wigler, 1987. Rigorous feedback control of cAMP levels in Saccharomyces cerevisiae. Genes Dev. 1: 931–937. [DOI] [PubMed] [Google Scholar]
  32. Nocero, M., T. Isshiki, M. Yamamoto and C. S. Hoffman, 1994. Glucose repression of fbp1 transcription of Schizosaccharomyces pombe is partially regulated by adenylate cyclase activation by a G protein α subunit encoded by gpa2 (git8). Genetics 138: 39–45. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Ohi, R., A. Feoktistova and K. L. Gould, 1996. Construction of vectors and a genomic library for use with his3-deficient strains of Schizosaccharomyces pombe. Gene 174: 315–318. [DOI] [PubMed] [Google Scholar]
  34. Pearson, B. M., Y. Hernando and M. Schweizer, 1998. Construction of PCR-ligated long flanking homology cassettes for use in the functional analysis of six unknown open reading frames from the left and right arms of Saccharomyces cerevisiae chromosome XV. Yeast 14: 391–399. [DOI] [PubMed] [Google Scholar]
  35. Shiozaki, K., and P. Russell, 1995. Cell-cycle control linked to extracellular environment by MAP kinase pathway in fission yeast. Nature 378: 739–743. [DOI] [PubMed] [Google Scholar]
  36. Shiozaki, K., and P. Russell, 1996. Conjugation, meiosis, and the osmotic stress response are regulated by Spc1 kinase through Atf1 transcription factor in fission yeast. Genes Dev. 10: 2276–2288. [DOI] [PubMed] [Google Scholar]
  37. Stettler, S., E. Warbrick, S. Prochnik, S. Mackie and P. Fantes, 1996. The wis1 signal transduction pathway is required for expression of cAMP-repressed genes in fission yeast. J. Cell Sci. 109: 1927–1935. [DOI] [PubMed] [Google Scholar]
  38. Stiefel, J., L. Wang, D. A. Kelly, R. T. K. Janoo, J. Seitz et al., 2004. Suppressors of an adenylate cyclase deletion in the fission yeast Schizosaccharomyces pombe. Eukaryot. Cell 3: 610–619. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Sugiura, R., T. Toda, H. Shuntoh, M. Yanagida and T. Kuno, 1998. pmp1+, a suppressor of calcineurin deficiency, encodes a novel MAP kinase phosphatase in fission yeast. EMBO J. 17: 140–148. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Sugiura, R., T. Toda, S. Dhut, H. Shuntoh and T. Kuno, 1999. The MAPK kinase Pek1 acts as a phosphorylation-dependent molecular switch. Nature 399: 479–483. [DOI] [PubMed] [Google Scholar]
  41. Toda, T., I. Uno, T. Ishikawa, S. Powers, T. Kataoka et al., 1985. In yeast, RAS proteins are controlling elements of adenylate cyclase. Cell 40: 27–36. [DOI] [PubMed] [Google Scholar]
  42. Watanabe, Y., Y. Lino, K. Furuhata, C. Shimoda and M. Yamamoto, 1988. The S. pombe mei2 gene encoding a crucial molecule for commitment to meiosis is under the regulation of cAMP. EMBO J. 7: 761–767. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Welton, R. M., and C. S. Hoffman, 2000. Glucose monitoring in fission yeast via the gpa2 Gα, the git5 Gβ, and the git3 putative glucose receptor. Genetics 156: 513–521. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Xue, Y., M. Batlle and J. P. Hirsch, 1998. GPR1 encodes a putative G protein-coupled receptor that associates with the Gpa2p Gα subunit and functions in a Ras-independent pathway. EMBO J. 17: 1996–2007. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Yun, C. W., H. Tamaki, R. Nakayama, K. Yamamoto and H. Kumagai, 1998. Gpr1p, a putative G-protein coupled receptor, regulates glucose-dependent cellular cAMP level in yeast Saccharomyces cerevisiae. Biochem. Biophys. Res. Commun. 252: 29–33. [DOI] [PubMed] [Google Scholar]

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