Present address: College of Agriculture and Life Sciences, Seoul National University, Seoul, Korea.
A putative G protein-coupled receptor negatively controls sexual development in Aspergillus nidulans
Article first published online: 4 FEB 2004
Volume 51, Issue 5, pages 1333–1345, March 2004
How to Cite
Han, K.-H., Seo, J.-A. and Yu, J.-H. (2004), A putative G protein-coupled receptor negatively controls sexual development in Aspergillus nidulans. Molecular Microbiology, 51: 1333–1345. doi: 10.1111/j.1365-2958.2003.03940.x
- Issue published online: 4 FEB 2004
- Article first published online: 4 FEB 2004
- Accepted 14 November, 2003.
G protein-coupled receptors (GPCRs) are key components of heterotrimeric G protein-mediated signalling pathways that detect environmental signals and confer rapid cellular responses. To broaden our understanding of signalling mechanisms in the filamentous fungus Aspergillus nidulans, intensive analyses of the Aspergillus nidulans genome have been carried out and nine genes (gprA∼gprI) that are predicted to encode seven transmembrane spanning GPCRs have been identified. Six of nine putative GPCRs have been disrupted and the gprD gene was found to play a central role in coordinating hyphal growth and sexual development. Deletion of gprD (ΔgprD) causes extremely restricted hyphal growth, delayed conidial germination and uncontrolled activation of sexual development resulting in a small colony covered by sexual fruiting bodies. Genetic studies indicate that GprD may not signal through the FadA (Gα)-protein kinase A (PKA) pathway. Elimination of sexual development rescues both growth and developmental abnormalities caused by ΔgprD, suggesting that the primary role of GprD is to negatively regulate sexual development. This is supported by the fact that environmental conditions inhibiting sexual development suppress growth defects of the ΔgprD mutant. We propose that the GprD-mediated signalling cascade negatively regulates sexual development, which is required for proper proliferation of A. nidulans.
Heterotrimeric G protein-mediated signalling pathways play a central role in transmembrane signalling in eukaryotes by modulating extracellular signals as they are transmitted into the cell. The basic unit of heterotrimeric G protein signalling is comprised of three parts, a seven-transmembrane-spanning G protein coupled receptor (GPCR), a heterotrimeric G protein consisting of α, β and γ subunits, and an effector, such as an enzyme or ion channel. When bound to their ligands, GPCRs are sensitized in an active conformation, which increases affinity of heterotrimeric G proteins to GPCRs. Binding of heterotrimeric G proteins to GPCRs causes conformational changes of their associated Gα subunits and such a Gα conformational change results in GDP-GTP exchange of Gα subunits. GTP-bound Gα subunits undergo another conformational switch that promotes the dissociation of Gα-GTP and Gβγ subunits, and both Gα-GTP and Gβγ can amplify and propagate signals by modulating activities of one or more effector molecules (for reviews see Lengeler et al., 2000; Neves et al., 2002).
In mammals, GPCRs and G protein components mediate responses to light, flavour, odours, numerous hormones, neurotransmitters and other signals (for reviews see Baldwin, 1994; Strader et al., 1994). In fungi, heterotrimeric G proteins mediate responses to pheromones or nutrients (e.g. glucose or nitrogen starvation) and play essential roles in cell growth, mating, cell division, cell-cell fusion, morphogenesis and chemotaxis. Moreover, pathogenic fungi rely on external signals that allow the fungi to recognize their hosts (for review see Bölker, 1998; Versele et al., 2001). Roles of heterotrimeric G proteins in host-sensing and pathogenesis in Cryptococcus neoformans (Lengeler et al., 2000), Magnaporthe grisea (Liu and Dean, 1997), Cryphonectaria parasitica (Choi et al., 1995), and Ustilago maydis (Regenfelder et al., 1997) have been reported.
In the model filamentous fungus Aspergillus nidulans, a heterotrimeric G protein and its proper regulation play a central role in maintaining the genetically programmed lifecycle represented by hyphal growth and development. Hyphal growth signalling is mediated by both the α (FadA) and β (SfaD) subunits of a heterotrimeric G protein (Yu et al., 1996; Rosén et al., 1999). When FadA (Gα) is in its active-GTP bound-state, it is dissociated with its cognate SfaD:Gγ (a hypothetical Gβγ dimer) and the separated FadA and SfaD:Gγ can both activate downstream effectors for proliferation. Constitutive activation of FadA growth signalling results in fluffy-autolytic phenotype caused by uncontrolled proliferation of hyphal mass. Mutations leading to such elevated hyphal growth phenotype also cause a blockage in sexual/asexual development and production of the mycotoxin sterigmatocystin (ST: Yu et al., 1996; Hicks et al., 1997; Adams and Yu, 1998; Rosén et al., 1999). Activation of asexual and sexual development requires that this FadA-SfaD:Gγ signalling is, at least partially, inactivated and FlbA plays a key role in this growth inhibition. FlbA is an RGS (regulator of G protein signalling) domain protein that rapidly turns off growth signalling, likely by acting as GTPase activating protein (GAP) on FadA (Lee and Adams, 1994a; Yu et al., 1996; Hicks et al., 1997). Loss of flbA function results in fluffy-autolytic colonies that lack sexual/asexual sporulation caused by constitutive activation of FadA-signalling. Recent studies show that FadA growth signalling is transduced in part via protein kinase A (PKA; Shimizu and Keller, 2001).
Compared to the recent progress with G proteins and downstream effectors, despite their importance, no GPCRs have been studied in A. nidulans. To broaden our understanding of G protein-mediated signalling mechanisms controlling hyphal growth and various aspects of development, we decided to identify and characterize all putative GPCRs in A. nidulans. Analyses of filamentous fungal genomes have been carried out employing various GPCRs, and nine genes that are predicted to encode putative GPCRs with seven transmembrane (hepta-helices) spanning domains have been identified and designated as gprA–gprI for G protein (coupled) receptor. Six of the nine putative GPCR-encoding genes have been disrupted. Whereas disruption of gprC, gprE or gprG did not cause clearly distinguishable phenotypic changes, deletion of gprA and gprB, similar to yeast pheromone sensing receptor genes, resulted in abnormal sexual development (unpubl. data). Most importantly, we found that GprD, somewhat similar to the yeast glucose sensing Gpr1p (Kraakman et al., 1999; Lorenzo et al., 2000), plays a key role in coordinating hyphal growth and sexual development in A. nidulans. Further genetic analyses of gprD indicate that GprD is required to negatively modulate sexual development, which in turn confers proper hyphal growth and balanced development. GprD does not appear to act through the FadA-PKA signalling pathway, which negatively controls asexual development. Here we present characterization and genetic analyses of gprD as well as a newly proposed model for signalling mechanisms controlling hyphal growth and sexual development.
Identification of nine genes predicted to encode putative GPCRs in the A. nidulans genome
In order to identify all putative GPCRs in Aspergillus nidulans we searched the following filamentous fungal genome databases (DB): A. nidulans DB provided by Monsanto (AnDB; http://microbial.cereon.com/) and Whitehead Institute Center for Genome Research (WICGR: http://www.genome.wi.mit.edu/annotation/fungi/aspergillus/index.html), Aspergillus fumigatus DB from TIGR (AfDB; http://www.tigr.org/tdb/e2k1/afu1/), and Neurospora crassa DB from WICGR (NcDB; http://www.genome.wi.mit.edu/annotation/fungi/neurospora/). Whereas GPCRs respond to variety of ligands, they all contain a conserved structure of seven transmembrane (7-TM) spanning domains. The following GPCRs were utilized for searches; Saccharomyces cerevisiaeα-factor receptor Ste2p (Burkholder and Hartwell, 1985), a-factor receptor Ste3p (Hagen et al., 1986), glucose-sensing Gpr1p (Xue et al., 1998; Kraakman et al., 1999; Lorenzo et al., 2000), Schizosaccharomyces pombe glucose-sensing Git3p (Welton and Hoffman, 2000), nitrogen starvation-sensing Stm1p (Chung et al., 2001), pheromone receptor Mam2p (Kitamura and Shimoda, 1991) and Map3p (Tanaka et al., 1993), and Dictyostelium discoideum cAMP receptor cAR1 (Klein et al., 1988). Combinatorial searches of fungal genomes resulted in the identification of nine genes that are predicted to encode GPCRs with 7-TM domains (hepta-helices) in AnDB (Table 1). These putative GPCR genes are designated as gprA – gprI (G protein-coupled receptor A–I).
|Genes and linkage groupa||Protein used for tblastn (organism)b||Pairwise blastpc||Gene coding and physical information|
|E-value||Identities||Positives||♯ of a.a (kDa)||Contig||ORF region in contig defined in this study/ coding strand/♯ of intrond||Locus♯ and coding regions predicted by WICGR/strand|
|gprAe (VII)||Ste2p||2e-21||80/285||140/285||377||1.43||2284–1083/–/1||AN2520.1 2519–1080/–|
|gprB e (IV)||Ste3p||5e-35||89/296||146/296||349||1.132||14491–13401/–/1||AN7743.1 14491–12943/–|
|gprC (II)||Git3p/Gpr1pf||9e-04||41/226||77/226||439||1.61||692136–690772/–/1||AN3765.1 692136–690769/–|
|(S. pombe/S. cerevisiae)||(18%)||(33%)||(48.9)|
|gprD (VI)||Git3p/Gpr1pf||0.81||33/213||78/213||427||1.55||393119–391742/–/1||AN3387.1 393119–391739/–|
|(S. pombe/S. cerevisiae)||(15%)||(36%)||(48.3)|
|gprE (IV)||GprD||e-136||267/482||321/482||493||1.170||81047–79496/–/1||AN9199.1 81047–79463/–|
|gprF (II)||Stm1p||4e-30||101/353||156/353||369||1.145||72232–71025/–/1||Not defined|
|gprG (VIII)||Stm1p||3e-17||82/327||130/327||377||1.16||181797–180526/–/0||Not defined|
|gprH (II)||cAR1p||5e-06||32/114||54/114||404||1.145||76385–77741/+/3 h||AN8262.1h 76385–77744/–|
|gprI (NDg)||GprH||1e-36||114/363||170/363||328||1.151||22608–21448/–/3||AN8348.1 22213–23242/–|
Phylogenetic analyses of nine putative A. nidulans GPCRs with other fungal GPCRs group them into four classes (Fig. 1): (i) GprA and GprB similar to the pheromone-sensing Ste2p and Ste3p in S. cerevisiae, respectively; (ii) GprC, GprD, and GprE similar to glucose sensing Gpr1p in S. cerevisiae; (iii) GprF and GprG similar to Stm1p in S. pombe; (iv) GprH and GprI similar to cAMP receptors, cAR1p found in D. discoideum (references described above). Six of nine GPCRs have been disrupted and the most dramatic changes resulted from the deletion of the gprD gene. In this study, further characterization of the gprD gene is presented. Characterization and functional analyses of other putative receptor genes will be published elsewhere.
Gene structure and mRNA levels of gprD
The gene structure of gprD, i.e. 1378 bp ORF with a 98 bp intron (Fig. 2A) was determined by comparison of sequences of genomic DNA and the RT-PCR product of gprD. The gprD gene is predicted to encode a 427 amino acid polypeptide with 7-TM (hepta-helical) domains. Multiple alignment of S. pombe Git3p with GprC, GprD and GprE shows high similarity among these GPCRs, particularly within 7-TM regions (shown by green underlines in Fig. 2B). To ensure that gprD encodes a real (expressed) gene, steady-state mRNA levels of gprD throughout the lifecycle of A. nidulans were examined by Northern blot analysis. As shown in Fig. 3A, the gprD gene is found to encode an 1.5 kb transcript that accumulates in conidia, growing vegetative hyphae (submerged culture), and early phase of asexual development (air exposed solid culture). The gprD mRNA levels decrease after 16 h post asexual induction. Interestingly, no gprD mRNA is detectable after 12 h post sexual induction (air exposed with sealing as described in Han et al., 2001; 2003).
Deletion of gprD results in restricted hyphal growth and uncontrolled activation of sexual development
To characterize function of GprD, the deletion mutant (ΔgprD) was generated. A disruption construct, where the gprD ORF was replaced by the argB+ selectable marker with ∼ 1.0 kb gprD flanking regions, was made by our newly developed DJ-PCR tool (see Experimental procedures). This disruption construct was transformed into the recipient host strain PW1 (biA1; argB2; methG1) and transformants were screened for the ΔgprD pattern by genomic DNA isolation followed by PCR. Random screening of 30 transformants resulted in the isolation of one ΔgprD mutant. We then meiotically crossed this putative ΔgprD mutant with various developmentally wild-type strains and confirmed that phenotypic characteristics observed in the ΔgprD mutant were solely because of the absence of gprD. Phenotypes of the ΔgprD mutant presented below are a summary of examination of the multiple ΔgprD mutants.
Deletion of gprD causes pleiotropic effects. As shown in Fig. 3B, the ΔgprD mutant shows extremely restricted hyphal growth at 37°C on minimal medium (MM) compared to the wild type. Whereas typical wild-type strains reached about 50 mm diameter at 4 days post inoculation, the ΔgprD mutant essentially shows no hyphal growth at 37°C on MM. Rather, it initially forms undifferentiated aerial hyphae and then produces dense Hülle cell aggregates with brief conidiophore development in the centre of the colony (Fig. 3C). It appears that radial extension of the ΔgprD mutant is achieved by forming Hülle cells. This phenotype was more obvious when the colonies were formed from a single conidium than point inoculation (data not shown). The ΔgprD mutant then produced an excessive number of fruiting bodies (cleistothecia) covering the whole colony at 14 days at 37°C MM without any forced induction of sexual development (Fig. 3C). It is important to note that under the same culture conditions wild-type strains (FGSC26 or other isogenic strains) form hardly any cleistothecia. Collectively, with the assumption that deletion of gprD eliminated associated downstream signalling, GprD is predicted to mediate signalling that is necessary for activating growth and/or inhibiting sexual development.
GprD is required for proper germination
We microscopically examined conidial germination and hyphal tip growth characteristics of the ΔgprD mutant on solid MM. As shown in Fig. 4, although deletion of gprD did not affect swelling of conidia, it caused an approximately 3 h delay of germination and limited hyphal tip growth compared to wild type. Similar results were obtained when these strains were grown in liquid MM submerged culture (data not shown). These results indicate that GprD is required for both proper germination and hyphal growth. We then examined patterns of septum formation and nuclei distribution in the ΔgprD mutant by staining its hyphae with calcofluor and DAPI, respectively, and found no abnormalities relative to the extension (growth) of the hyphae (data not shown), indicating that the growth defect of the ΔgprD mutant is not the result of improperly coordinated cell cycle progression or cytokinesis.
GprD-mediated signalling is likely independent to FadA-PkaA
Because gprD encodes a putative GPCR that is apparently required for hyphal growth, we examined the likelihood of GprD being an upstream receptor of FadA and SfaD:Gγ (see Introduction). First, we compared phenotypes of the ΔgprD and fadAG203R (dominant interfering FadA mutation; Yu et al., 1996) mutants. We expected that if GprD specifically mediates signalling to activate hyphal growth through FadA (and thus SfaD:Gγ), then ΔgprD would cause lack of both FadA and cognate SfaD:Gγ signalling resulting in the similar phenotype to fadAG203R. The fadAG203R mutation is predicted to block dissociation of FadA-GTP from SfaD:Gγ and it causes restricted hyphal growth, highly elevated asexual development but lack of sexual development. These phenotypic changes are almost opposite to the ones caused by ΔgprD especially in terms of developmental patterns, suggesting that FadA is likely not the cognate Gα for GprD. Second, we generated and examined the ΔgprD; ΔflbA double mutant. Previously, we showed that loss-of-function or dominant interfering mutations in fadA or sfaD could suppress fluffy-autolytic phenotype caused by ΔflbA (Yu et al., 1996; Rosén et al., 1999). If gprD is the receptor for the FadA signalling pathway, deletion of gprD should cause the absence (or inhibition) of activation (GDP/GTP exchange) of FadA (Gα) thereby resulting in the suppression of ΔflbA. However, as shown in Fig. 5A, the double mutant ΔgprD; ΔflbA is indistinguishable from the ΔflbA mutant, suggesting that GprD is likely not the GPCR necessary for GDP/GTP exchange of FadA. Finally, the ΔgprD; ΔpkaA double mutant was generated and examined. Compared to ΔgprD or ΔpkaA single mutant, the ΔgprD; ΔpkaA double mutant conidiates to a level in between two single mutants and shows non-conidiating aerial hyphae at the growing edge of the colony on MM at 37°C (Fig. 5B and C). Collectively, these results indicate that GprD-mediated signalling is independent and parallel to the FadA-PkaA pathway with possible cross-talking (see Discussion).
Primary role of GprD is to negatively control sexual development
As described above and presented in Fig. 3C, in addition to restricted hyphal growth, the ΔgprD mutant shows uncontrolled activation of sexual development. Two hypotheses regarding the primary role of the GprD-dependent signalling cascade can be derived from these results: (i) GprD mediates signalling necessary for activating growth, which in turn represses sexual development, or (ii) GprD-mediated signalling negatively regulates sexual development that confers proper hyphal growth.
To further dissect the role of GprD in controlling sexual development and to correlate molecular events with phenotypic changes caused by the absence of gprD, mRNA levels of the gene encoding a sexual specific transcription factor (NsdD) were examined. As shown in Fig. 6A, nsdD mRNA accumulated at much higher levels in the ΔgprD mutant than in wild type both during vegetative growth (submerged) and asexual development (air exposed), suggesting that the absence of gprD gene results in de-repression of nsdD gene expression.
As nsdD or veA are required for normal fruiting body formation, loss-of-function mutations in either nsdD or veA result in lack of sexual development (Han et al., 2001; Kim et al., 2002). To test our hypothesis on the primary role of GprD, we generated double mutants between ΔgprD and ΔnsdD or ΔveA. If the GprD signalling cascade's main role is to activate hyphal growth, elimination of sexual specific functions in ΔgprD would cause small colonies without sexual development. However, deletion of nsdD or veA suppresses growth defect of ΔgprD to the single mutant level without sexual development (Fig. 6B), suggesting that the primary role of GprD-signalling is to negatively control sexual development rather than activating growth and the limited hyphal growth caused by ΔgprD is likely caused by constitutive and uncontrolled activation of sexual development.
Environmental conditions favouring asexual development suppress growth and developmental defects caused by ΔgprD
One of the unique characteristics of the ΔgprD mutant was that growth and developmental defects (uncontrolled sexual development) were rescued by the presence of salt (0.6 M KCl or 0.8 M NaCl), sorbitol (1.2 M) or yeast extract (YE; see Fig. 7A). Moreover, when the ΔgprD mutant was cultured at room temperature from the start, it did not show any signs of growth defects or developmental abnormality relative to wild type (Fig. 7B). Salt-, sorbitol-, or low temperature- mediated growth recovery can be explained, at least in part, by the fact that these conditions inhibit sexual development, which in turn confer growth recovery of the ΔgprD mutant. For the effects of YE, one can speculate that addition of YE provides substances to the ΔgprD mutant which enhanced growth, but not necessarily asexual development.
We also examined the effects of acetate (a poor carbon source) on ΔgprD. In A. nidulans, acetate promotes asexual development while inhibiting formation of cleistothecia (Han et al., 2003). Consistently, the ΔgprD mutant grown on MM with acetate as a sole carbon source formed no cleistothecia and the mutant colonies were about 70% size of wild type (not shown). Taken together, these results support our model that the primary role of GprD is to negatively control sexual development, which in turn confers proper hyphal growth.
G protein-coupled receptors (GPCRs) form one of the largest protein families in humans. This might be the case for the filamentous fungal species, too. Our findings of at least nine genes with seven transmembrane domains (α-helices) in the A. nidulans genome and a report of 10 putative GPCR genes in N. crassa (Galagan et al., 2003) support the fact that GPCRs might well form one of the largest protein families in filamentous fungi. We have also analysed the A. fumigatus genome and found seven GPCRs closely related to A. nidulans GPCRs (not shown). By similarity analysis, nine A. nidulans GPCRs are grouped into four, where two or three are grouped together, suggesting possible redundant functions. Whereas three putative GPCRs, GprC, GprD, and GprE, are grouped together with the yeast glucose sensing Gpr1p, yeast Gpr1p is apparently evolutionarily distant from these proteins. GprC, GprD and GprE appear to be unique in filamentous fungi and share high levels of similarity (only) within the group. Deletion of gprC or gprE apparently causes no detectable phenotypes, suggesting that these two putative GPCRs may share redundant functions. Redundancy of GPCR functions will make it difficult to precisely dissect specific signalling mediated by an individual GPCR and its associated ligand(s).
Complexity of heterotrimeric G protein signalling is even further enhanced when we account for other primary components functioning in transmitting signals. For instance, like other filamentous fungal species, A. nidulans has three Gα subunits, one Gβ subunit and one Gγ subunit (designated as GpgA; unpubl. results). If we assume that all nine putative GPCRs have the ability to activate certain G protein signalling, one Gα might be activated by more than one GPCR. Thus, if this is the case, elimination of one GPCR function may not result in lack of downstream signalling. Moreover, other components that can bind to GPCRs and modulate GPCR-mediated signalling need to be counted. It is known that GPCRs can bind to various modulators including phospholipase C (PLC; Ansari et al., 1999) and ankyrin repeat-containing protein (Givan and Sprague, 1997) as well as heterotrimeric G proteins. Furthermore, recent studies showed that certain GPCRs might mediate signalling independent of heterotrimeric G proteins, and, thus, it has been suggested that GPCRs be called hepta-helical receptors (Hall et al., 1999). Taken together, one cannot draw solid conclusions by examining phenotypic changes resulting from a disruption of a single GPCR.
Although a simplified model for the antagonistic activities of hyphal growth and asexual development has been proposed (Yu et al., 1996; Rosén et al., 1999), regulation of sporulation by FadA-mediated signalling is rather complicated. Although it is clear that both FadA and SfaD positively mediate hyphal growth, primary roles of individual subunits in developmental regulation appear to be different. The absence (or interference) of SfaD (Gβ) activity, either by ΔsfaD or fadAG203R(dominant interfering mutation), led to extremely reduced growth yet uncontrolled activation of asexual sporulation, indicating that SfaD-mediated growth signalling is required for proper control of conidiation. However, these fadA or sfaD mutants could not sporulate without an early developmental activator called FluG (Lee and Adams, 1994b; Yu et al., 1996), indicating that control of conidiation by FadA-SfaD signalling is indirect. Moreover, elimination of asexual sporulation by deleting fluG does not rescue growth defects of ΔsfaD or fadAG203R (Yu et al., 1996; Rosén et al., 1999), suggesting that two signalling pathways are independent and parallel to each other and the primary role of the FadA-SfaD (and PKA; Shimizu and Keller, 2001) signalling cascade is to activate proliferation, which in turn represses asexual sporulation.
Our findings in this study suggest that, like the antagonistic relationship between asexual sporulation and FadA-PkaA growth signalling, there is a specific GPCR-mediated signalling cascade that coordinates sexual development (fruiting body formation) versus growth. This is supported by the facts that: (i) mRNA accumulation of a critical sexual development specific transcription factor, NsdD, is highly upregulated even in early vegetative growth phase and asexually induced conditions; (ii) elimination of sexual development by deleting nsdD or veA suppresses growth and developmental defects of the ΔgprD mutant; and (iii) environmental factors such as poor carbon source, high osmolarity, and low temperature that favour asexual development over sexual development rescue growth defects of ΔgprD. The antagonistic relationship between sexual development and hyphal growth was further tested by generating multiple nsdD overexpression strains (in collaboration with D.-M. Han's group at Wonkwang University, Korea), where nsdD is fused under the alcA promoter. It has been shown that overexpression of asexual development functions inhibits growth and even germination of spores (Adams et al., 1988; Mirabito et al., 1989; Adams and Timberlake, 1990). Likewise, the alcA(p)::nsdD mutant showed restricted growth during induced condition (data not shown), confirming that sexual development and growth are antagonistic to each other and that coordinated control of sexual development is necessary for proper hyphal growth. However, unlike ΔgprD, overexpression of nsdD did not lead to the formation of cleistothecia, indicating that the absence of gprD activity might upregulate multiple components necessary for completion of sexual development.
What does GprD likely activate? The fact that no Gα deletion or dominant interfering mutants (Yu et al., 1996; K.-Y. Jahng, pers. comm.) show similar phenotypes to ΔgprD makes it hard to speculate about G proteins associated with GprD (if any). Furthermore, limited genetic/biochemical data along with the complexity of development in A. nidulans do not allow us to even speculate what would be ligand(s) for GprD. Thus, we will only discuss possible downstream effector components. In general, heterotrimeric G proteins activate PKA, PKC (protein kinase C) and/or MAPK (mitogen-activated protein kinase), which control a broad range of targets. In an attempt to identify a downstream signalling cascade activated by GprD, we briefly examined the effects of the absence of gprD on PKA and PKC activity (not shown). Wild type showed both PKA and PKC activity and the ΔpkaA mutant showed undetectable PKA with wild-type level PKC activity. However, no PKC and very low PKA activity was detected in the ΔgprD mutant grown in liquid MM. Interestingly, addition of YE restored PKA but not PKC activity in the ΔgprD mutant, implying that GprD is required for both PKC and PKA activity and requirement of GprD for PKA activity can be bypassed by the addition of YE. Taken together, it can be speculated that GprD mediates signalling primarily through PKC, yet can affect PKA activity (cross-talking; Fig. 8).
In mammalian systems, PLC activates PKC through the generation of membrane-bound diacylglycerol (DAG) and increased cytosolic Ca++ concentration (for review see Rhee and Bae, 1997). However, in yeast, neither Ca++ nor DAG, but small GTP-binding proteins (e.g. Rho1) activate PKC (for review see Perez and Calonge, 2002), with possible cross-talking between the PLC and PKC pathways. Phenotypic alterations by various environmental conditions allow us to further speculate on the GprD signalling branch based on previous studies in yeast S. cerevisiae reporting that: (i) budding yeast contains a single PLC gene (PLC1); (ii) Plc1p-deficient mutants arrest at temperatures above 35°C as multibudded, enlarged cells unable to complete cytokinesis; (iii) growth of Plc1p-deficient mutants is recovered when the mutants are grown at 25°C; and (iv) addition of sorbitol (5%) into YPD restored yeast growth above 35°C (Yoko-o et al., 1993; Ansari et al., 1999). These characteristics are somewhat consistent with those of the ΔgprD mutant. In addition, recent studies showed that PLC directly interacts with Gpa2 (Gα) and Gpr1p (GPCR) in yeast, suggesting the positive role of PLC in a growth control pathway (Ansari et al., 1999). A gene (plcA) encoding a probable PLC has been identified and possible interactions among GprD, PLC and PKC remain to be investigated.
Fungal strains, growth conditions and genetic manipulations
Strains used in this study are listed in Table 2. Minimal medium (MM) with appropriate supplements was prepared following standard procedures (Käfer, 1977). If needed, yeast extract was added (0.1% or 0.5% final concentration). All strains were cultured at 37°C unless otherwise indicated. Standard genetic and transformation techniques were employed (Pontecorvo et al., 1953; Yelton et al., 1984). The gprD deletion strain TKH02.66 was generated by transforming the recipient strain, PW1, with the PCR-generated deletion construct. TKH02.66 was meiotically crossed with RJYE07, TKIS18.11, KHH32 and DVAR1 to generate RKH62.4, RKH65.3, RKH67.13 and RKH64.14 respectively. In most experiments, the ΔgprD mutant was grown in MM with 0.5% YE and 0.6 M KCl to facilitate growth and conidiation and, conidia were collected (see Text).
|PW1||biA1; argB2; methG1; veA1||P. Weglenski|
|TKH02.66||biA1; argB2; methG1; ΔgprD::argB +; veA1||This study|
|RKH57.25||biA1; argB–b; ΔgprD::argB +; veA1||This study|
|TKIS18.11||pabaA1, yA2, ΔpkaA::argB +; ΔargB::trpC +; trpC801, veA1||Shimizu and Keller (2001)|
|RKH65.3||pabaA1, yA2, ΔpkaA::argB +; argB –b; ΔgprD::argB +; veA1||This study|
|RJYE07||biA1, ΔflbA::argB+; ΔfadA::argB +, trpC801, veA1||Hicks et al. (1997)|
|RKH62.3||biA1, ΔflbA::argB +; methG1; veA1||This study|
|RKH62.4||biA1, ΔflbA::argB +methG1; ΔgprD::argB+; veA1||This study|
|KKH32||pabaA1, yA2; ΔargB::trpC +ΔnsdD::argB+; trpC801, veA1||Han et al. (2001)|
|RKH67.13||pabaA1, yA2; argB –b; ΔnsdD::argB +; ΔgprD::argB +; veA1||This study|
|DVAR1||pabaA1, yA2; ΔargB::trpC +; trpC801, ΔveA::argB +||Kim et al. (2002)|
|RKH64.14||pabaA1, yA2; argB–b; ΔgprD::argB +; ΔveA::argB +||This study|
Nucleic acids manipulations
Genomic DNA extraction was carried out as previously described (Seo et al., 2003). Briefly, mycelial mats obtained from liquid stationary cultures (∼18 h at 37°C) were dried by squeezing with paper towels and transferred to microcentrifuge tubes containing 0.5 mm zirconia/silica bead (BioSpec Products, OK). Samples were ground by a mini bead-beater (BioSpec Products, OK) with equal volumes of breaking buffer (2% Triton X-100, 1% SDS, 0.1 M NaCl, 10 mM Tris-Cl pH 8.0, 1 mM EDTA) and phenol:chloroform:isoamyl alcohol (25 : 24 : 1). The aqueous phase was separated by centrifugation, and genomic DNA was harvested by ethanol precipitation. For total RNA isolation, 0.1–0.2 g of mycelia were ground by bead-beating in the presence of 1 ml Trizol reagent (Invitrogen, CA). RNA extraction from the ground hyphae was performed as manufacturer's instruction. Northern blot analysis was carried out as previously described (Yu et al., 1996; Hicks et al., 1997). MagnaProbe membranes (Osmonics, MN) were used for blotting the nucleic acids after separation of 15 µg of total RNA in 1.1% agarose gels containing 3% formaldehyde. 32P-labelled probes were used to hybridize using modified Church buffer (Hicks et al., 1997) at 63°C for 16–20 h. Polymerase chain reaction products amplified by using primers OKH122 – OKH127 and OKH188 – OKH189 (Table 3) were used as gprD and nsdD gene-specific probes, respectively. For producing a PCR-based deletion construct of the gprD gene, double-joint PCR (DJ-PCR) method was employed, which is similar to the method described in Davison et al. (2002). Briefly, primers were designed for flanking region amplification (∼1.0 kb) of the gprD gene. Primers away from the gprD ORF carried ∼25 bases of homo-logous sequence overlapping with the ends of the selectable marker, argB+. Three (5′-and 3′ flanking regions and argB+) amplicons were mixed as 1 : 2 : 1 ratio and the second round of thermo-cycling was carried out. Using a new nested primer pair, the resulting product of the second round PCR reaction was amplified. The final deletion construct was composed of ∼1.0 kb flanking regions and deleted gprD gene region that was replaced with the argB+ marker. This final PCR product was directly used for transforming PW1.
|Name||Sequence (5′-to 3′)||Description|
|OKH60||GACTCTATACCACCGTACGCCGATAT||Forward primer for argB +|
|OKH61||CACCGGGTGCGATTTGCCCCATTTCC||Reverse primer for argB +|
|OKH099||GGATACCGTCAACTAAGCCAAT||gprD deletion 5′-forward|
|OKH100||AGTCAAATGAGGCCTCTAAACTGGTCAGAT GCGGCGTTGTAGAGTTTAG||gprD deletion 5′-reverse with complementary 5′- argB+ tail (bold)|
|OKH101||AGCCAAGGTAGATCCAGGCCTAACACATCG TGATGATTCTACCGAGGAG||gprD deletion 3′- forward with complementary 3′- argB+ tail (bold)|
|OKH102||GTCAGGGACCGCTAGACACAAG||gprD deletion 3′- reverse|
|OKH103||GGAGTAGACGCAGCCTGG||gprD deletion construct 5′-nest|
|OKH104||TGAGAGAGATGGGGGAGG||gprD deletion construct 3′-nest|
|OKH122||TGCACTGGCCGTTTCAC||gprD ORF forward primer for probe amplification|
|OKH127||GACCGGTCGTAGTGAAG||gprD ORF reverse primer for probe amplification|
|OKH188||CTCGCACTGTCAAATCAA||nsdD ORF forward primer for probe amplification|
|OKH189||GGCTCCAGCTTCCAGAAC||nsdD ORF reverse primer for probe amplification|
PKA and PKC activity assay
Samples were obtained by inoculating two loopfuls of conidia (∼ 1.0 × 107) to 2 ml of liquid MM with or without 0.5% YE in test tubes, which were slanted 45° and incubated at 37°C for 20 h without shaking (stationary liquid culture). Individual mycelial mats were collected, lyophilized and ground. Ground samples were mixed with 500 µl of extraction buffer (25 mM Tris-HCl pH 7.4, 10 mM MgCl2, 150 mM NaCl, 1 mM DTT, 1 mM EDTA). After incubation on ice for 30 min, samples were centrifuged at 10 000 g for 10 min at 4°C. The supernatant was collected and protein concentration was measured with BCA protein assay reagent kit (Pierce, IL) according to the manufacturer's instruction. Measurements of PKA and PKC activity were carried out using SignaTECT cAMP-dependent protein kinase (in the presence of cAMP) and SignaTECT protein kinase C assay systems (Promega, WI) respectively.
Microscopy and photographs
Microscopic images were taken by using an Olympus BH2 compound microscope with the Kodak MDS290 system. Culture plate photographs were taken using a SONY DSC-F707 digital camera.
We are thankful to Nancy Keller, Dong-Min Han and Kwang-Yeop Jahng for sharing fungal strains and unpublished information and Monsanto Company, the Whitehead Institute Center for Genome Research and TIGR for the genome databases. Special thanks go to Byron Brehm-Stecher and Ellin Doyle in our institute for critically reviewing this manuscript. This work was supported by sponsors of the Food Research Institute, a Hatch from the College of Agricultural and Life Sciences and funds from Graduate School at the University of Wisconsin-Madison to J.H.Y.
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