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Chitin is an essential component of the fungal cell wall and its synthesis is under tight spatial and temporal regulation. The fungal human pathogen Candida albicans has a four member chitin synthase gene family comprising of CHS1 (class II), CHS2 (class I), CHS3 (class IV) and CHS8 (class I). LacZ reporters were fused to each CHS promoter to examine the transcriptional regulation of chitin synthesis. Each CHS promoter had a unique regulatory profile and responded to the addition of cell wall damaging agents, to mutations in specific CHS genes and exogenous Ca2+. The regulation of both CHS gene expression and chitin synthesis was co-ordinated by the PKC, HOG MAP kinase and Ca2+/calcineurin signalling pathways. Activation of these pathways also resulted in increased chitin synthase activity in vitro and elevated cell wall chitin content. Combinations of treatments that activated multiple pathways resulted in synergistic increases in CHS expression and in cell wall chitin content. Therefore, at least three pathways co-ordinately regulate chitin synthesis and activation of chitin synthesis operates at both transcriptional and post-transcriptional levels.
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The fungal cell wall is a dynamic structure whose composition and structural organization is regulated during the cell cycle and in response to changing environmental conditions, imposed stresses and mutations in cell wall biosynthetic processes (reviewed in Klis et al., 2006; Ruiz-Herrera et al., 2006). Chitin and β(1–3)-d-glucan, represent the main structural components of the fungal cell wall. These polysaccharides oppose the positive turgor pressure within the cell and ultimately determine the morphology of the cell (Munro and Gow, 2001; Klis et al., 2002; Roncero, 2002). Chitin and glucan synthesis therefore play fundamental roles in maintaining fungal cell integrity during growth and morphogenesis and in adaptation to stress (Cabib, 1987; Wessels, 1990; Shaw et al., 1991; Sietsma and Wessels, 1994; Gooday, 1995). Because these structural polysaccharides do not occur in mammals and are essential for fungi, there is considerable potential for cell wall synthesis as a target for antifungal drugs (Munro and Gow, 1995; Munro et al., 2001; Odds et al., 2003). New generation echinocandins that target the synthesis of cell wall β(1–3)-d-glucan are proving effective agents in the treatment of opportunistic fungal pathogens such as Candida albicans (Denning, 2003). Chitin synthase inhibitors have not yet been discovered that have clinical use in the treatment of fungal infections (Odds et al., 2003).
Regulation of chitin synthesis occurs both at the transcriptional and post-translational levels and is dependent on precise targeting and activation of chitin synthases to specific locations in the plasma membrane, and the provision of adequate substrate (Munro and Gow, 1995). All fungi examined to date have multiple genes encoding chitin synthase families (Munro and Gow, 2001; Roncero, 2002; Ruiz-Herrera and San-Blas, 2003). Individual chitin synthase enzymes perform distinct functions at specific stages of the cell cycle. Saccharomyces cerevisiae has three chitin synthase enzymes – Chs1p (Class I), Chs2p (Class II) and Chs3p (Class IV) while C. albicans has four chitin synthases – two class I enzymes –CaChs2p and CaChs8p, CaChs3p (Class IV) and CaChs1p (a class II enzyme which is the orthologue of ScChs2p). Relatively little is known about the transcriptional regulation of chitin synthase genes in fungi but considerable attention has been focused on post-transcriptional regulation by Chs4–7, which influences Chs3p chitin synthase activation and localization in S. cerevisiae and C. albicans. ScChs7p controls exit of ScChs3p from the ER, ScChs5p and ScChs6p regulate its exit from the trans-Golgi network (Ziman et al., 1996; Santos and Snyder, 1997; Santos et al., 1997; Ziman et al., 1998; Trilla et al., 1999). ScChs4p tethers ScChs3p to the septins at the mother-bud neck via ScBni4p (Demarini et al., 1997; Trilla et al., 1997). Chitin synthesis is therefore influenced by endogenous and exogenous factors that directly and indirectly regulate the chitin synthase catalytic proteins.
Disruption of genes in cell wall biosynthetic pathways of S. cerevisiae and C. albicans often results in alteration and redistribution of chitin and β(1–3)-d-glucan in the cell wall, the synthesis of new cell wall proteins and changes in their cross-linking to cell wall polysaccharides (reviewed in Popolo et al., 2001; Klis et al., 2002; Klis et al., 2006). Defects in cell wall integrity are sensed by the transmembrane proteins of the Mid2p and the Wscp family, which signal via the Rom2p guanine nucleotide exchanger leading to activation of the Rho1p GTPase. Rho1p has many downstream targets including protein kinase C and the β(1–3)-d-glucan synthase subunits Fks1p and Fks2p (Popolo et al., 2001). In S. cerevisiae this ‘cell wall salvage’ or ‘cell wall compensatory’ pathway is activated in response to cell wall perturbing agents such as Calcofluor white (CFW), Congo Red (CR), caffeine, β-glucanases and cell wall mutations and is mediated primarily through the PKC cell integrity MAP kinase cascade and its downstream target the transcription factor Rlm1p (Lagorce et al., 2003; Boorsma et al., 2004; Garcia et al., 2004). In S. cerevisiae, elevation of chitin levels in response to activation of the salvage pathway is largely dependent upon ScChs3p (Valdivieso et al., 2000; Carotti et al., 2002). Several studies have highlighted the importance of signalling systems in co-ordinating this regulation. A higher proportion of ScChs3p localized to the plasma membrane in heat-stressed cells (Valdivia and Schekman, 2003). This mobilization of ScChs3p was dependent upon activation of Rho1p and Pkc1p, and the phosphorylation of ScChs3p by Pkc1p.
A second MAP kinase cascade, the high osmolarity glycerol response (HOG) pathway, has also been suggested to play a role in regulating cell wall architecture in S. cerevisiae (Garcia-Rodriguez et al., 2000; Kapteyn et al., 2001) and in C. albicans (Eisman et al., 2006). In S. cerevisiae, the HOG pathway is required for the response to CFW and mutants in several components of the pathway are resistant to CFW (Garcia-Rodriguez et al., 2000). In addition, changes in osmotic pressure have been shown to regulate chitin synthase activity in the dimorphic fungus Benjaminiella poitrasii suggesting the HOG pathway is involved in chitin regulation (Deshpande et al., 1997).
Transcript profiling studies have implicated Ca2+ in the regulation of ScCHS1 (Yoshimoto et al., 2002). In addition, sequences recognized by the Ca2+/calcineurin-dependent transcription factor Crz1p/Tcn1p have been identified upstream of a number of genes that are upregulated in cell wall mutants that activate the cell wall salvage pathway (Lagorce et al., 2003; Boorsma et al., 2004; Garcia et al., 2004; Karababa et al., 2006). These studies directed us towards examining the role of Ca2+ signalling in the regulation of chitin synthesis in C. albicans.
Each of the four C. albicans Chs enzymes plays a distinct role in cellular growth. CaChs1p synthesizes the septal chitin and contributes to chitin in the lateral cell wall and is essential for viability in both the yeast and hyphal forms (Munro et al., 2001). CaChs2p encodes the major chitin synthase activity in vitro, and chs2Δ null mutants have fractionally less chitin in hyphal cells (Gow et al., 1994; Munro et al., 1998). CaChs3p synthesizes the majority of the chitin in the lateral cell wall and the ring of chitin at the site where a new bud emerges (Bulawa et al., 1995; Mio et al., 1996). CaChs8p and CaChs2p account for almost all the measurable in vitro chitin synthase activity in membrane preparations but are non-essential for growth (Munro et al., 2003). In C. albicans, northern analyses suggested that CaCHS2 and CaCHS3 are upregulated shortly after induction of hyphal formation while CaCHS1 is expressed at low but constant levels in both yeast and hyphae (Chen-Wu et al., 1992; Munro et al., 1998). Hyphal formation in C. albicans is accompanied by a three to fivefold increase in the chitin content of the cell wall (Chattaway et al., 1968; Sullivan et al., 1983; Munro et al., 1998).
Here we examine the regulation of chitin synthesis of C. albicans and describe the signalling pathways that co-ordinate this process. We used a lacZ reporter gene fused to the putative promoters of each of the C. albicans CHS genes to test hypotheses about the expression of CHS genes when cells are challenged with cell wall perturbing agents or subjected to environmental stresses. We show that transcriptional regulation of the CHS genes is stimulated via at least three pathways – the PKC and HOG MAP kinase cascades and the Ca2+/calcineurin pathway. Each of the four chitin synthase promoters was regulated differentially, but all were activated by exogenous Ca2+ in a calcineurin and Crz1p-dependent manner. In addition, hyper-stimulation of CHS gene expression was observed when multiple signalling pathways were activated simultaneously and this resulted in greatly elevated cell wall chitin levels.
Endogenous CHS promoter activity in wild-type cells and chsΔ mutants
Transcriptional activity of the four chitin synthase genes of C. albicans was characterized using a lacZ reporter system. Plasmid placpoly 6 containing URA3 and RPS1 was used to create a fusion between the promoter of each C. albicans chitin synthase gene and the Streptococcus thermophilus lacZ open reading frame (ORF). A 1 kb region upstream from the ATG start codon of CHS1, CHS2, CHS3 and CHS8 was cloned into placpoly 6 generating, respectively, plasmids pCHS1plac, pCHS2plac, pCHS3plac and pCHS8plac. Ura–C. albicans cells were transformed with each linearized plasmid, homologous recombination resulted in integration of the plasmid at the RPS1 locus and Ura+ transformants were selected. Transformants were screened by Southern blot analysis and those with a single copy integration of pCHSplac (strains NGY210-NGY213, Table 1) were analysed further. The CHS2 and CHS3 promoters had the highest and lowest level of expression, respectively, for growth in YPD medium (P < 0.05) (Fig. 1). Real-time quantitative polymerase chain reaction (PCR) confirmed these results (data not shown).
Table 1. Candida albicans strains used in this study.
The pCHSplac plasmids were transformed into isogenic mutant strains derived from CAI-4 with single or double CHS gene disruptions (Table 1) to test whether deletion of CHS genes results in a compensatory upregulation of other members of the CHS family. The CHS1 promoter activity was significantly increased in the single mutants chs3Δ (strain Myco3) and chs8Δ (NGY138) and the double mutants chs2Δchs8Δ (NGY128) and chs2Δchs3Δ (C157) (Fig. 2). Chs1p may contribute to the maintenance of lateral wall integrity (Munro et al., 2001) and play a compensatory role when CHS3 and CHS8 gene functions are lost. Expression from the CHS8 promoter was stimulated when either CHS2 or CHS3 were deleted and was increased further in the chs2Δchs3Δ double null mutant. CHS3 expression was slightly elevated in chs8Δ and chs2Δchs8Δ mutants while the CHS2 promoter did not show any significant changes in any of the mutants tested (Fig. 2). Therefore, the deletion of single CHS genes resulted in activation of the expression of others.
CHS promoter activity responds to wall perturbing agents
Transcriptional regulation of the four CHS genes was determined in response to various environmental changes and perturbations (Fig. 3). Growth at 37°C stimulated CHS1, CHS2 and CHS3 promoters compared with growth at 25°C (not shown) and 30°C (control conditions). The addition of SDS, which perturbs membrane integrity, or CFW that interferes with cell wall assembly, induced expression from three of the four promoters (Fig. 3). CR stimulated only CHS1 expression. Caffeine, an inhibitor of cAMP phosphodiesterase, stimulates dual phosphorylation of ScSlt2, the MAP kinase component of the PKC cell wall integrity signal transduction pathway (Martin et al., 2000). The addition of 12 mM caffeine to the growth medium resulted in significantly elevated expression from all four CHS promoters.
The CHS transcriptional response to cations and salts – 200 mM Ca2+, Mn2+, K2+, Li2+, Mg2+ or 800 mM NaCl was tested. Addition of K2+, Li2+, Mg2+ or Na2+ had no detectable effects (data not shown), however, exogenous Ca2+ and Mn2+ activated all four CHS promoters (Fig. 4). Some response was observed even with 5 mM Ca2+ (data not shown). Exogenously applied Ca2+ leads to activation of the calcineurin pathway, which induces dephosphorylation of the Crz1p transcription factor (Cyert, 2003). The calcineurin specific inhibitors FK506 and Cyclosporin A inhibited Ca2+ activation of CHS transcription (Fig. 4). Activation of CHS expression with Mn2+ was also reduced, but not totally blocked, by simultaneous addition of FK506 (data not shown) suggesting the Mn2+-specific activation occurred in part through the calcineurin signalling pathway but also involved a calcineurin-independent mechanism. These results suggest that Ca2+ and Mn2+ activated CHS expression via both calcineurin/Crz1-dependent and independent mechanisms. The calmodulin inhibitor chloropromazine had no effect on CHS transcriptional activity but chloropromazine with 200 mM Ca2+ completely inhibited the Ca2+-activation response. The calcium ionophore A23187 also inhibited the Ca2+-activation response (data not shown). Inhibition by chloropromazine suggested that the observed Ca2+ stimulation involved the classical Ca2+ signalling pathway acting through calmodulin and calcineurin.
To corroborate these findings the four CHS-reporter constructs were transformed into null mutant strains lacking genes involved in the calcineurin pathway. The Crz1p transcription factor is dephosphorylated when the phosphatase calcineurin is activated by Ca2+/calmodulin. It then enters the nucleus and induces expression of a number of genes, many of which encode proteins with cell wall-related functions (Yoshimoto et al., 2002; Lagorce et al., 2003; Garcia et al., 2004; Karababa et al., 2006; Pardini et al., 2006). Putative calcium-dependent response element (CDRE) motifs that are recognized by the Crz1p transcription factor were found in the promoter region of the CHS genes in C. albicans (Table 2). In the cna1Δ strain, which is mutated in one of the calcineurin subunits, the CHS2 and CHS8 promoters still responded to Ca2+, but the response of the CHS2 promoter was reduced (Fig. 5). The Ca2+ responses of the CHS1 and CHS3 promoters were not significantly different to the untreated cna1Δ control. Stimulation with Ca2+ was abrogated by simultaneous addition of FK506. In the crz1Δ mutant background, expression from the CHS1 promoter was elevated when cells were grown in YPD, and addition of Ca2+ did not further stimulate CHS1 expression (Fig. 5). The response of the CHS2 and CHS3 promoters to exogenous Ca2+was significantly different in the crz1Δ mutant compared with wild-type cells. Therefore, CHS2 and CHS3 were activated in part via the Ca2+/calcineurin/Crz1 pathway but Crz1p repressed the expression of CHS1. Transformation of the CHSp-lacZ constructs into the double calcium channel mutant mid1Δcch1Δ had no significant effect on Ca2+-stimulated activation of expression (data not shown).
Synergistic stimulation of CHS promoters by combined Ca2+ and CFW treatment
Addition of Ca2+ or CFW stimulated the CHS promoters – therefore we tested the effects of combinations of Ca2+ and CFW treatments. All four promoters were hyper-stimulated by combined treatment with Ca2+ and CFW (Table 3). The pCHSplac plasmids were transformed into mutant strains lacking MAP kinase genes of the PKC and HOG signal transduction pathways (Table 1). The first mutant tested had a disrupted MKC1 gene that encodes the MAP kinase of the PKC pathway (Navarro-Garcia et al., 1998). In the mkc1Δ strain background the CHS8 promoter, and to a lesser extent, the CHS2 promoter had reduced activity. All four CHS promoters were still stimulated by Ca2+ in the mkc1Δ mutant (Table 3), therefore the Ca2+-induced upregulation of CHS promoters can be independent of the PKC pathway. However, the ability of the CHS2 and CHS8 promoters to respond to CFW was impaired in the mkc1Δ mutant. All four promoters were stimulated by the combined Ca2+/CFW treatment but the level of stimulation of CHS2 and CHS8 promoters was significantly less in mkc1Δ cells compared with wild-type cells. The response of the CHS promoters to CFW and combined Ca2+/CFW was also examined in the crz1Δ mutant (Table 3). Again the response of CHS2 and CHS8 promoters to CFW was significantly reduced in the crz1Δ mutant and the Ca2+/CFW induction of all four promoters was dramatically reduced (three- to five-fold).
Table 3. The CHS promoters are hyper-stimulated by combined Ca2+/CFW treatment.
Fold change with respect to the control in the same genetic background.
Ratio of fold change in the mutant compared with fold change in wild-type cells under the same conditions.
Statistically significant changes are highlighted in bold.
100 ± 7
174 ± 38
190 ± 34
888 ± 59
91 ± 9
189 ± 25
141 ± 41
735 ± 40
209 ± 11
202 ± 22
154 ± 19
411 ± 174
39 ± 12
48 ± 9
47 ± 13
592 ± 219
100 ± 3
226 ± 43
221 ± 23
889 ± 121
68 ± 3
196 ± 13
76 ± 3
379 ± 18
142 ± 10
153 ± 12
147 ± 6
402 ± 43
43 ± 3
54 ± 2
38 ± 1
354 ± 113
100 ± 7
154 ± 26
118 ± 10
693 ± 33
101 ± 8
184 ± 23
104 ± 25
732 ± 93
110 ± 8
89 ± 6
91 ± 4
288 ± 41
156 ± 14
251 ± 21
135 ± 9
988 ± 237
100 ± 9
158 ± 16
263 ± 14
1237 ± 421
27 ± 5
76 ± 17
16 ± 2
378 ± 47
141 ± 8
160 ± 12
233 ± 16
529 ± 94
43 ± 8
48 ± 5
34 ± 6
315 ± 69
Three of the four CHS promoter sequences contained ATF/CREB elements – potential binding sites for the Sko1p transcription factor that is regulated by Hog1p (Table 2) (Proft et al., 2005). Expression from the CHS1, CHS2 and CHS8 promoters was reduced in the hog1Δ mutant compared with wild-type cells in YPD and in the presence of Ca2+ or CFW (Table 3). In contrast, expression from the CHS3 promoter was increased in the hog1Δ mutant, suggesting Hog1p normally repressed CHS3 transcription. In the hog1Δ mutant the CHS3 promoter still responded to exogenous Ca2+, but not to CFW and combinations of Ca2+ and CFW stimulated the CHS3 promoter in both the hog1Δ and wild-type backgrounds. The Ca2+/CFW-induced stimulation of the CHS1 and CHS2 promoters was not significantly altered in the hog1Δ strain but the level of expression from the CHS8 promoters was significantly less than in wild-type cells. Together these data suggest that Mkc1p, Crz1p and Hog1p play significant roles in the Ca2+/CFW hyper-stimulation of CHS promoters.
We tested whether the Ca2+-activated CHS gene expression translated into measurably higher chitin synthase enzyme activity. C. albicans yeast cells were cultured in YPD or YPD plus 100 mM Ca2+ for 5 h and membrane fractions of wild type (CAI-4), chsΔ and signalling mutant strains were prepared and assayed for chitin synthase activity. The specific chitin synthase activity of wild-type mixed membrane fractions (MMF) increased slightly when exogenous Ca2+ was added to the growth medium (Fig. 6). In the mkc1Δ mutant, Chs activity was comparable to the control strain and did not increase upon addition of Ca2+. The hog1Δ mutant had markedly elevated Chs activity compared with the control and addition of Ca2+ did not further stimulate chitin synthase activity. Chitin synthase activity of the crz1Δ mutant was comparable to wild type and decreased in Ca2+-treated cells. As shown previously (Munro et al., 2003), the Chs activity of the chs2Δchs8Δ double mutant was only around 5% of wild-type levels and no further stimulation was observed when cells were grown in the presence of Ca2+. The chs3Δ mutant had reduced Chs activity but this was elevated in response to Ca2+. Therefore, Chs2p and Chs8p are mainly responsible for the elevated Chs activity in response to Ca2+ and this was mediated via Crz1p, Mkc1p and Hog1p. Attempts were made to measure chitin synthase activity from membranes prepared from cells grown in the presence of CFW and Ca2+/CFW. No detectable Chs activity was found (not shown). CFW inhibition of in vitro chitin synthase activity has been reported previously (Roncero and Duran, 1985) and was shown to be dependent upon pH of the growth medium (Roncero et al., 1988).
Treatment with Ca2+ and CFW elevates cell wall chitin levels
The chitin content of cells was measured under conditions where Ca2+ stimulated CHS gene expression and in vitro Chs activity. Addition of Ca2+, CFW and Ca2+/CFW resulted in elevated cell wall chitin levels in wild-type CAI-4 cells (Fig. 7) with the combination of Ca2+/CFW giving the greatest stimulation. Chs3p is responsible for the synthesis of the majority of the chitin in the C. albicans cell wall (Bulawa et al., 1995; Mio et al., 1996). In the chs3Δ mutant, chitin levels were dramatically reduced and Ca2+-treatment had only a slight effect on chitin content. The chs2Δchs8Δ mutant behaved similarly to wild type. In the mkc1Δ mutant, chitin levels were lower than in parental controls, but were elevated after combined Ca2+/CFW treatment. However, addition of Ca2+ or CFW alone had little effect on chitin content in the mkc1Δ mutant. Chitin levels of untreated crz1Δ cells were significantly higher than wild-type cells again suggesting that under some conditions Crz1p represses chitin synthesis. Chitin levels of crz1Δ were elevated by CFW or Ca2+/CFW treatments but did not respond, or were repressed, when treated with Ca2+ alone. Untreated hog1Δ cells had wild-type chitin levels that were increased marginally when cells were grown with Ca2+ but the activation with CFW or Ca2+ plus CFW was significantly reduced compared with wild-type cells. These results suggest that elevated cell wall chitin content in response to combined treatments with Ca2+ and CFW is due mainly to Chs3p and that the PKC and HOG and to a lesser extent the Ca2+/Crz1 pathways are involved in this Chs3p-dependent stimulation of chitin synthesis.
Ca2+- and CFW-dependent phosphorylation of Mkc1p and Cek1p
The phosphorylation status of the Mkc1p and Cek1p MAP kinases was examined in order to assess the status of the PKC and SVG (STE vegetative growth) pathways in the strains and treatments described above (Navarro-Garcia et al., 2005; Eisman et al., 2006). Cek1p is the C. albicans orthologue of ScKss1p, which is the S. cerevisiae MAP kinase component of pathways that regulate filamentous growth, the pheromone response and promote vegetative growth (the latter via the SVG pathway) (Lee and Elion, 1999; Eisman et al., 2006). The SVG pathway is constitutively activated in the och1ΔN-glycosylation mutants of S. cerevisiae and C. albicans (Lee and Elion, 1999; Bates et al., 2006) and in the hog1Δ mutant and has been implicated in the response to cell wall perturbing agents (Eisman et al., 2006). Phospho-Mkc1p and phospho-Cek1p were identified by western analysis as 59 kDa and 48 kDa bands, respectively, using phospho-specific antibodies (Fig. 8). Phosphorylation status was assessed 10, 30, 60 and 120 min after treatment addition, however, only 10 and 120 min time points are presented here. In non-stressed conditions, no phospho-Mkc1p was detected in wild type or hog1Δ yeast cells however, Mkc1p was phosphorylated in the crz1Δ mutant. Activation of Mkc1p was observed in the crz1Δ strain after treatment with Ca2+. CFW stimulated strong activation of Mkc1p in the wild type, weaker activation in hog1Δ and activation comparable to untreated controls in crz1Δ. CFW-stimulated phosphorylation of Mkc1p was observed at 2 h in wild type and hog1Δ and after 1 h in crz1Δ. Combined treatment with Ca2+ and CFW had a synergistic effect on activation of Mkc1p in the crz1Δ strain where phospho-Mkc1p was detected after 10 min. In wild-type cells, Ca2+/CFW did not give as strong a response in terms of Mkc1p phosphorylation as CFW alone. In agreement with Navarro-Garcia et al. (2005); Roman et al. (2005) and Eisman et al. (2006), phospho-Mkc1p was only detected in extracts prepared from hog1Δ cells when cells were treated with CFW. Under these conditions Cek1p appeared to be constitutively activated in the hog1Δ mutant. Treatment of the hog1Δ strain with Ca2+, CFW and Ca2+/CFW increased the level of phospho-Cek1p significantly. We conclude that the PKC pathway is activated when cells are treated with CFW and Ca2+/CFW but Ca2+ alone could not stimulate phosphorylation of Mkc1p in a wild-type background.
This study has shown that at least three signalling systems are involved in chitin synthesis regulation: (i) Ca2+/calcineurin/Crz1p, (ii) PKC-Mkc1p and (iii) HOG pathways. At the transcriptional level CHS expression was monitored using a lacZ reporter gene fused to each of the four C. albicans CHS promoters. Each promoter was regulated differentially – the CHS2 promoter was the most active under control conditions (YPD at 30°C) and the CHS3 promoter was the least active. Real-time quantitative PCR confirmed these observations (data not shown). The CHS promoters responded to deletion of other CHS genes with a twofold increase in expression levels from CHS1, CHS3 and CHS8 promoters in several chsΔ mutants. In Wangiella dermatitidis, a melanized fungal pathogen of humans, a compensatory increase in WdCHS expression has also been described in response to chitin synthase gene disruptions (Wang et al., 2002). Although there is no evidence of true functional redundancy within the chitin synthases examined to date, fungi appear to upregulate certain CHS in compensation for loss of others perhaps to maintain a robust cell wall.
The C. albicans CHS promoters were found to respond to a number of environmental stimuli notably when cells were treated with cell wall perturbing drugs and when growth medium was supplemented with Ca2+. Addition of exogenous Ca2+, stimulated CHS gene expression; stimulated in vitro chitin synthase activity, and resulted in increased cell wall chitin mediated through Chs3p. In addition, simultaneous treatment of cells with CFW and Ca2+ resulted in synergistically enhanced expression from all four CHS promoters and a threefold increase in the amount of chitin in the cell wall.
The CHS promoters were activated by exogenous Ca2+ and Mn2+ but not by equivalent concentrations of Mg2+ or Na+. In S. cerevisiae, Ca2+ activates calcineurin via calmodulin, which induces gene expression by regulating the Crz1p/Tcn1p transcription factor. This plays a role in regulating cell wall structure including the induction of ScCHS1 in response to Ca2+ (Yoshimoto et al., 2002) and tolerance of fungi to a wide range of antifungal agents (Edlind et al., 2002; Sanglard et al., 2003; Onyewu et al., 2004; Karababa et al., 2006). Ca2+-activation of CaCHS expression was blocked by inhibitors of both calmodulin and calcineurin confirming the role of this pathway in the regulation of CHS genes. In addition, transcription from all CHS promoters was reduced in the cna1Δ mutant in response to exogenous Ca2+. In the crz1Δ mutant, basal activity of the CHS2, CHS3 and CHS8 promoters was not altered but the CHS1 promoter was de-repressed. In addition, crz1Δ cells were attenuated, but not completely blocked, in their ability to activate CHS expression in response to exogenous Ca2+ and the hyper-stimulation of CHS expression caused by cross-activation with Ca2+ and CFW was reduced dramatically in the crz1Δ mutant background. In silico analysis of the CHS promoter sequences also identified potential CDREs, motifs recognized by Crz1p. Therefore, activation of the CHS promoters due to Ca2+ was mainly regulated by the classical Cna1/Crz1 pathway; however, some of the Ca2+ stimulation was Cna1p and/or Crz1p-independent indicating that calcineurin and Crz1p may have roles that are distinct from their role in this signalling pathway. Our results suggest that in C. albicans the Ca2+ signalling pathway plays a major role in regulating chitin synthesis. This pathway may be vital to the co-ordination of responses to a variety of conditions that compromise cell wall integrity because it also regulates the expression of genes encoding cell wall proteins Utr2p and Crh11p (Pardini et al., 2006) and the glucan synthase catalytic subunit Fks1p/Gsc1p (Sanglard et al., 2003).
Many of the conditions that stimulated the CHS promoters including growth at 37°C, treatment with cell wall perturbing agents CFW, CR and SDS and the cAMP-phosphodiesterase inhibitor caffeine lead to hyper-phosphorylation of Slt2p/Mkc1p (De Nobel et al., 2000; Martin et al., 2000; Navarro-Garcia et al., 2005). We used the mkc1Δ MAP kinase null mutant to test the role of the PKC pathway in chitin synthesis regulation. In the mkc1Δ mutant background the CHS2 and CHS8 promoters were less responsive to CFW but were still stimulated by Ca2+ and compared with wild-type cells only the CHS2 promoter had a significantly reduced response to Ca2+/CFW. However, there was a dramatic decrease in chitin levels in the mkc1Δ mutant under all conditions tested suggesting post-transcriptional regulation of Chs3p occurs via the PKC pathway. This pathway has been shown to regulate Chs3p in S. cerevisiae (Valdivia and Schekman, 2003).
The HOG signalling pathway was the third pathway implicated in CHS transcriptional regulation. Promoter sequences recognized by the HOG-regulated transcription factor Sko1p were identified in the CHS1, CHS3 and CHS8 promoters. Loss of the HOG pathway in non-stressed conditions resulted in reduced expression of CHS1, CHS2 and CHS8 but increased expression of CHS3. Therefore, as with Crz1p, blocking a particular signalling pathway had both positive and negative effects on CHS expression. Although the CHS1, CHS2 and CHS8 promoters had attenuated responses to either CFW or Ca2+ in the hog1Δ mutant, only the CHS8 promoter had markedly reduced activity in response to the combined Ca2+/CFW treatment compared with wild-type cells. Despite the apparently low basal level of expression of CHS2 and CHS8 in the hog1Δ mutant, the levels of chitin synthase enzyme activity were greater than in wild-type cells. This suggests the presence of a compensatory mechanism that is activated in response to loss of Hog1p that acts post-transcriptionally and results in enhanced Chs enzyme activity. Nevertheless, the amount of chitin in the wall of the hog1Δ mutant synthesized in response to co-stimulation with Ca2+ and CFW was significantly (40%) lower than wild type implying that the HOG pathway is involved in activation of chitin synthesis via Chs3p.
The Ca2+/calcineurin, PKC and HOG pathways contribute to the hyper-stimulation of chitin synthesis in response to Ca2+/CFW treatment. The role of the Ca2+-signalling pathway appears to be mainly in regulating CHS transcription, whereas the PKC and HOG pathways also contribute to regulation of chitin synthase enzyme activity and total cell wall chitin content. The Mkc1 pathway positively regulates CHS expression, chitin synthase activity and chitin levels in the cell wall while the Ca2+/Crz1 and HOG pathways have both positive and negative regulatory effects on different CHS genes and Chs isoenzymes (Fig. 9).
Interpretation of experiments studying CHS expression is complicated by cross-talk between signalling pathways and by compensatory mechanisms that are triggered in mutants defective in single signalling pathways. For example, mutants blocked in the HOG pathway have a constitutively active Cek1 MAP kinase, which contributes to a CR resistance phenotype (Roman et al., 2005; Eisman et al., 2006). We examined the phosphorylation status of Mkc1p and Cek1p in cells treated with Ca2+, CFW and Ca2+/CFW. We confirmed phosphorylation of Cek1p in the hog1Δ mutant and enhanced phosphorylation of Cek1p when hog1Δ was treated with Ca2+, CFW and Ca2+/CFW. Despite activation of Cek1p, chitin levels are reduced in the hog1Δ mutant suggesting Cek1p may not make a major contribution to chitin regulation. Our findings also corroborate the observations of Navarro-Garcia et al. (2005) that Hog1p was required for phosphorylation of Mkc1p under a variety of conditions, but not with CFW treatment. In addition, our Western analyses suggested that Mkc1p was phosphorylated in the crz1Δ mutant. These data suggest that these pathways do not operate in isolation and that mutations in one pathway results in activation of others as in the case of Cek1p activation in the hog1Δ mutant and Mkc1p phosphorylation in the crz1Δmutant. Hence, the activation of Mkc1p is not solely responsible for the elevated chitin synthesis under the conditions tested. Instead, the PKC, HOG and Ca2+ signalling pathways all contribute to the regulation of chitin synthesis.
In conclusion, the Ca2+/Crz1p, PKC-Mkc1p and HOG signalling pathways co-ordinate the regulation of chitin synthesis in C. albicans. The use of multiple pathways may enable the fungus to fine-tune the co-ordinated assembly of cell wall chitin to exogenous stresses by modulating chitin synthesis. CHS gene expression responded to a wide range of environmental conditions and individual CHS genes and Chs enzymes responded differently to these stresses. This regulation is vital for the maintenance of a robust cell wall during growth and morphogenesis but also under conditions where the integrity of the cell wall is compromised by treatments with antifungal drugs that target fungal cell wall synthesis.
Strains, media and growth conditions
Candida albicans strains used in this study are listed in Table 1. C. albicans cultures were maintained on solid YPD medium comprising 1% (w/v) yeast extract, 2% (w/v) peptone, 2% (w/v) glucose, 2% (w/v) agar. Yeast cells of C. albicans were grown at 30°C in YPD with shaking at 200 r.p.m.
Transformation of C. albicans
Ura–C. albicans strains were cultured in 10 ml of YPD supplemented with 25 μg ml−1 uridine at 30°C for 36–72 h. After centrifugation, the cell pellets from 200 μl of cells were resuspended in 100 μl of OSB (200 mM LiAc pH 7.5, 100 mM DTT, 50% v/w PEG 6000, 10 mg ml−1 Clontech herring testis carrier DNA) and then transforming DNA was added. Samples were incubated at 43.5°C for 60 min and spread over SD agar plates (2% (w/v) d-glucose, 0.67% (w/v) yeast nitrogen base (YNB) (Bio 101, Carlsbad), 1.5% (w/v) purified agar, Oxoid) and incubated at 30°C. Single colonies were picked and grown in 5 ml of SD medium, and genomic DNA was extracted for Southern analysis.
Construction of plasmids and C. albicans strains
The placpoly-6 vector was used for the promoter-fusion reporter system and was based on a plasmid previously described by Uhl and Johnson (2001). This contains the CaURA3 and the CaRPS1 genes and was used to create fusion between the promoter of each CaCHS gene and the S. thermophilus lacZ ORF. A 1 kb upstream region from the ATG start codon of each CHS1, CHS2, CHS3 and CHS8 ORF was cloned into the PstI–XhoI sites of placpoly-6 generating pCHS1plac, pCHS2plac, pCHS3plac, pCHS8plac respectively. Ura–C. albicans cells were transformed with these plasmids previously cut within the RPS1 gene with StuI to target homologous recombination at the neutral chromosomal RPS1 locus and the URA3 gene was the selectable marker (Murad et al., 2000). Southern analysis was used to screen transformants and only those with single integrations of each pCHSplac plasmid were selected. Genomic DNA from each transformant was digested with XhoI/BamHI and hybridized to 693 bp RPS1 specific probe.
Measurement of β-galactosidase activity
The expression of each CHS gene in lacZ promoter fusions was measured using a modified version of the assays described previously (Rose and Botstein, 1983). C. albicans cells were grown with shaking at 200 r.p.m. at the chosen growth condition and harvested at OD600 < 1. Yeast cells were centrifuged at 3000 g for 5 min at 4°C and the pellet was resuspended in 0.5 ml of ice–cold water and transferred to microcentrifuge tubes. The cells were then centrifuged at 13 000 g for 5 min and resuspended in 0.5 ml of breaking buffer [100 mM TRIS-HCl pH 7.5, 0.01% (w/v) SDS, 1 mM dithiothreitol (DTT), 10% (v/v) glycerol, pepstatin 4 μg ml−1, 1 × proteinase cocktail tablets EDTA-free (Roche)]. Approximately equal volumes of glass beads (Sigma, Poole, UK G9268) and cell pellet were used and the cells were disrupted using a Fastprep cell breakage machine (Thermo Savant, Middlesex, UK) using six cycles of 30 s with chilling on ice for 1 min in between each cycle. The extract was centrifuged at 13 000 r.p.m. for 10 min and the protein concentration of the supernatant was measured using Coomassie® Protein Assay Reagent Kit (Pierce Biotechnology, Perbio, Rockford, UK). Varying quantities of protein extract, 30–300 μl, were added to Z-buffer (60 mM NaH2PO4, 40 mM Na2HPO4, 10 mM KCl, 1 mM MgSO4), in a total volume of 800 μl. The reaction was initiated by adding 200 μl of ο-nitrophenyl-β-d-galactopyranoside (ONPG) stock solution (4 mg ml−1 in phosphate buffer) and incubated until the yellow ο-nitrophenol product was produced. The reaction was stopped by addition of 400 μl of 1 M Na2CO3. The specific β-galactosidase activity was measured in terms of the yield of product ο-nitrophenol at the absorbance of 420 nm.
Measurement of chitin synthase activity
Mixed membrane fractions were prepared from exponential phase yeast cells and their chitin synthase activities measured as described previously (Munro et al., 1998). MMF proteins were activated by limited incubation with 100 ng trypsin μl−1 MMF at 30°C and the reactions were stopped by addition of 150 ng soybean trypsin inhibitor μl−1 MMF. Briefly, standard reactions for measuring chitin synthase activity were carried out in a 50 μl volume and were composed of; 50 μg MMF protein, 25 mM N-acetylglucosamine, 1 mM UDP-N-acetylglucosamine which included 25 nCi UDP-[U-14C]N-acetylglucosamine, 50 mM Tris-HCl pH 7.5 and 10 mM MgCl2. Incubations were carried out at 30°C for 30 min and the reaction was stopped by addition of 1 ml of 66% (v/v) ethanol. The reaction mixture was then filtered through GF/C filter discs (Whatman), which had been presoaked in 10% (v/v) trichloroacetic acid. The reactions tubes were rinsed out with 2 × 1 ml of 1% (v/v) Triton X-100 and each filter was then washed with 4 × 2 ml of 66% (v/v) ethanol. The radiolabelled chitin synthesized in the reaction was trapped on the filters and unincorporated substrate was removed by washing. Filters were dried at 80°C and their radioactivity counted in a scintillation counter.
Measurement of cell wall chitin content
Cell walls were prepared from 10 ml of C. albicans stationary phase yeast cultures grown in YPD and the chitin content was measured as described previously (Munro et al., 2003). Cells were disrupted with glass beads (Sigma, G9268) using a Fastprep cell breakage machine (Thermo Savant, Middlesex, UK) until at least 95% of cells were disrupted. They were then washed five times with 1 M NaCl and extracted in SDS-MerOH buffer (50 mM Tris, 2% sodium dodecyl sulphate (SDS), 0.3 M β-mercaptoethanol, 1 mM EDTA; pH 8.0) at 100°C for 10 min, then washed in dH2O. Cell wall pellets were resuspended in sterile dH2O, freeze dried, and the dry weight of recovered cell walls was measured. Chitin contents were determined by measuring the glucosamine released by acid hydrolysis of purified cell walls (Kapteyn et al., 2000).
Cells were grown in normal laboratory media and under conditions of various environmental stresses. Cells were grown overnight in YPD then transferred to YPD supplemented with different agents: 1 M sorbitol, 0.8 M NaCl, 0.2 M CaCl2, 100 μg ml−1 CFW, 200 μg ml−1 CR, 0.05% SDS, 12 mM caffeine, 25 mM DTT, 23 mM glucosamine, 50 μg ml−1 cyclosporin A, 1 μg ml−1 FK506, 1 mM chloropromazine, 4 μM A23187. Cells were harvested at OD600 0.8.
Western analysis was performed using the method of Millar et al. (1995) with some modifications. Overnight cultures of the wild type, crz1Δ and hog1Δ strains were diluted 1:50 into 25 ml of YPD supplemented with uridine and incubated shaking for 4 h at 30°C. The mid-log phase cultures were then treated with a final concentration of 100 mM CaCl2, 100 μg ml−1 CFW, or both for 0, 10, 30, 60 or 120 min. No-treatment controls were also performed. After treatment, cells were harvested by centrifugation (1500 g, 2 min, 4°C) and washed in 1 ml of cold lysis buffer (50 mM Tris-HCl pH 7.5, 150 mM NaCl, 0.5% NP40, 2 μg ml−1 Leupeptin, 2 μg ml−1 Pepstatin, 1 mM PMSF, 2 mM Na3VO4, 50 mM NaF). Cells were collected by centrifugation (800 g, 5 min, 4°C) and resuspended in 250 μl of cold lysis buffer. Cells were broken using a FastPrep machine in the presence of acid-washed glass beads (4 × 15 s bursts at speed 6.5 with 1 min on ice between bursts). The extracts were clarified by centrifugation (16 000 g, 5 min, 4°C). Protein concentration in the cleared lysate was estimated using the method described by Bradford (1976) with BSA as a standard.
Proteins were separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) using the XCell SureLockTM Mini-Cell system (Invitrogen) with NuPAGE®Novex Bis-Tris 4–12% precast gels (Invitrogen) in NuPAGE® MOPS-SDS Running Buffer (Invitrogen) containing NuPAGE® Antioxidant (Invitrogen) as per the manufacturer's instructions. Approximately 15 μg of protein was loaded in each lane. The proteins were transferred to InvitrolonTM PVDF Membranes (Invitrogen) in NuPAGE® Transfer Buffer containing methanol using the XCell IITM Blot Module (Invitrogen) following the manufacturer's instructions.
Following transfer, the membranes were rinsed in PBS and blocked in PBS-T + 10% BSA (PBS, 0.1% Tween-20, 10% (w/v) BSA, 50 mM (NaF) for 30 min at room temperature. The membranes were then incubated overnight at 4°C in PBS-T + 5% BSA (PBS, 0.1% Tween-20, 5% (w/v) BSA, 50 mM (NaF) containing a 1:1000 dilution of Phospho-p44/42 Map Kinase (Thr202/Tyr204) Antibody (Cell Signaling Technology). The membranes were washed five times for 5 min in PBS-T (PBS, 0.1% Tween-20) and then incubated for 1 h at room temperature in PBS-T + 5% BSA containing a 1:2000 dilution of Anti-rabbit IgG, HRP-linked Antibody (Cell Signaling Technology). The membranes were washed three times for 5 min in PBS-T and the signal was detected using LumiGLOTM Reagent and Peroxide (Cell Signaling Technology) as per the manufacturer's instructions.
Statistical significant differences in the assay results were determined with spss software using anova and Post Hoc Dunnett's T-test, P < 0.05. When the results displayed unequal variance the Kruskal–Wallis non-parametric test or Dunnett's T3 test were applied.
We thank D. Sanglard and J. Pla for provision of mutant strains and acknowledge financial support from the BBSRC (BB/C510191/1 and studentship to SS) the EC (Eurocellwall, Fungwall and SignalPath consortia), the Wellcome Trust (063204, 080088) and the MRC for a New Investigator Award to CAM. We are indebted to the referees for their valuable critiques.