In Candida glabrata, the sirtuins Sir2 and Hst1 control the expression of a wide number of genes including adhesins required for host colonization and niacin transporters needed for growth. Given that these sirtuins can be inactivated during infection, we asked if their inhibition could modify the response of C. glabrata to other stressful conditions. Here, we found that deletion of HST1 decreases susceptibility of C. glabrata to fluconazole and hydrogen peroxide. The transcription factor Pdr1 and the ABC transporter Cdr1 mediated the fluconazole resistance phenotype of the hst1Δ cells, whereas the transcriptional activator Msn4 and the catalase Cta1 are necessary to provide oxidative stress resistance. We show that the transcription factor Sum1 interacts with Hst1 and participate in the regulation of these genes. Interestingly, even though C. glabrata and Saccharomyces cerevisiae are closely related phylogenetically, deletion of HST1 decreased susceptibility to fluconazole and hydrogen peroxide only in C. glabrata but not in S. cerevisiae, indicating a different transcriptional control by two similar sirtuins. Our findings suggest that Hst1 acts as a regulator of stress resistance associated-genes.
Candida glabrata is a commensal yeast that acts as an opportunistic pathogen in immunocompromised patients. In the last three decades, C. glabrata has emerged as the second most frequent causative agent of mucosal and bloodstream fungal infections (Richardson and Lass-Florl, 2008; Pfaller and Diekema, 2010). Changes in the transcription program are a major strategy of C. glabrata to adapt to hostile environments. For example, induction of extracellular proteases, siderophore transporters, elements of the oxidative stress response (OSR), autophagy, shift in metabolism and chromatin remodelling are required for the survival of C. glabrata inside phagocytes (Kaur et al., 2007; Cuellar-Cruz et al., 2008; Roetzer et al., 2010; 2011; Nevitt and Thiele, 2011; Seider et al., 2011; Rai et al., 2012; Fukuda et al., 2013). In addition, C. glabrata induces membrane-spanning efflux pumps in the presence of fluconazole (Sanglard et al., 1999; Vermitsky and Edlind, 2004; Sanguinetti et al., 2005), which are associated with multidrug resistance (MDR) (Pfaller et al., 2010).
The response to environmental stimuli is essential not only to ensure the survival of C. glabrata within the host but also to detect conditions that favour infection. C. glabrata induces the transcription of virulence-related adhesins when availability of nicotinamide adenine dinucleotide (NAD+) precursors in the environment is limited (Domergue et al., 2005; Ma et al., 2009). This response is due to the derepression of epithelial adhesin genes encoded in subtelomeric regions that are subject to chromatin silencing (De Las Penas et al., 2003; Castano et al., 2005; Domergue et al., 2005). In C. glabrata, the Sir complex (Sir2, Sir3 and Sir4) is essential for subtelomeric silencing (De Las Penas et al., 2003; Castano et al., 2005; Rosas-Hernandez et al., 2008). The enzymatic component of the Sir complex is Sir2, a histone deacetylase that uses NAD+ as cofactor to keep the chromatin in a transcriptionally repressive state. Interestingly, since C. glabrata is unable to de novo synthesize NAD+ (Ma et al., 2007), Sir2 relies on the environmental supply of NAD+ precursors (niacin) to maintain silencing. Thus, environmental limitation of niacin causes inactivation of Sir2 and derepression of normally silent adhesins that result in increased adherence to epithelial cells (Domergue et al., 2005).
Sirtuins are a family of NAD+-dependent protein deacetylases present in all organisms, from bacteria to humans (Sauve et al., 2006). C. glabrata contains five sirtuin genes that include SIR2 and four homologues of SIR2 called HST1 – HST4. HST1 is the closest paralogue of SIR2 and both are involved in transcriptional repression, however these proteins display non-overlapping functions (Ma et al., 2009). In S. cerevisiae, the paralogues Sir2 and Hst1 have sub-functionalized after gene duplication (Mead et al., 2007; Froyd and Rusche, 2011) so that Sir2 operates through regional transcriptional silencing (Moazed et al., 1997; Tanny et al., 1999), whereas Hst1 acts as a local transcriptional repressor (Xie et al., 1999; McCord et al., 2003). The functional divergence between these sirtuins is likely conserved in C. glabrata (Domergue et al., 2005; Rosas-Hernandez et al., 2008; Ma et al., 2009). Hst1 controls the transcription of genes encoding for high-affinity niacin transporters (TNA1, TNR1 and TNR2) required to maintain NAD+ homeostasis in C. glabrata (Ma et al., 2009). When C. glabrata is starved for niacin, Hst1 losses its activity and the expression of these genes is derepressed, to ensure proper levels of NAD+ precursors that are required for growth (Ma et al., 2009). In S. cerevisiae, Hst1 also acts as a sensor that regulates the NAD+ levels, but mainly by controlling the biosynthetic NAD+ genes (Bedalov et al., 2003). It is worth noting that the transcriptional profile of the hst1Δ mutants between S. cerevisiae and C. glabrata is notably different (Hirao et al., 2003; Ma et al., 2009).
The auxotrophy of C. glabrata for NAD+ links the inactivation of sirtuins with the environment and the colonization of specific niches in the host, such as the urinary tract (Domergue et al., 2005; Ma et al., 2009). Given that sirtuins participate in transcriptional repression and can be inactivated in some niches of the host, we asked if the inactivation of sirtuins could modify the adaptive capacity of C. glabrata against stressful conditions. In this study, we decided to evaluate the role of the sirtuins in the OSR and the MDR of C. glabrata. The involvement of sirtuins in the control of either of the two processes remains largely unexplored. Here, we show the existence of a layer of epigenetic regulation that represses genes involved in the MDR and the OSR of C. glabrata. The induction of these responses is mediated by the specific inactivation of Hst1 and this control is present only in C. glabrata but not in the phylogenetically related yeast S. cerevisiae. Furthermore, Hst1 interacts with Sum1 and this interaction is required to repress its target genes in C. glabrata.
Inhibition of Hst1 decreases susceptibility to fluconazole in C. glabrata
In C. glabrata, sirtuins participate in the transcriptional regulation of a wide number of genes including the major virulence-associated adhesins (EPA genes) and transporters required for niacin assimilation (Domergue et al., 2005; Ma et al., 2009). Inactivation of sirtuins may occur in some niches of the host and can lead to the derepression of additional genes. We asked whether derepression of the remaining genes regulated by sirtuins affects the capacity of C. glabrata to adapt to other environmental stresses. We evaluated the involvement of sirtuins in two stress conditions: fluconazole exposure and oxidative stress. We spotted serial dilutions of stationary phase cells of the parental strain (BG14) on YPD plates in the absence or presence of 10 mM nicotinamide [NAM, a physiological non-competitive inhibitor of the NAD+-dependent histone deacetylases (Bitterman et al., 2002)] and different concentrations of fluconazole (see Experimental procedures). Interestingly, the parental strain (BG14) was able to grow at higher concentrations of fluconazole (32 μg ml−1) in the presence of NAM (Fig. 1A). This result indicated that sirtuins could be repressing genes involved in fluconazole resistance.
We then determined whether the sirtuins Sir2, Hst1 and Hst2, could participate in the resistance to fluconazole (see Experimental procedures). In principle, if one of these deacetylases controls fluconazole resistance genes, the deletion of the sirtuin will display a resistant phenotype. The sir2Δ and the hst2Δ mutant strains were able to grow at 8 μg ml−1 of fluconazole, the same concentration as the parental strain BG14 (Fig. 1A – NAM and data not shown). These results indicate that the reduced susceptibility to fluconazole observed in the presence of NAM, is not due to Sir2 or Hst2. In contrast, deletion of HST1 (hst1Δ) showed the same reduced susceptibility to fluconazole as the parental strain in the presence of NAM (Fig. 1A). The analysis of the single mutants, hst1Δ and sir2Δ treated with NAM, and the double mutant hst1Δ sir2Δ suggested that the reduced susceptibility to fluconazole is due to the relief of the Hst1-mediated repression (Figs 1A and S1). We then performed a quantitative fluconazole susceptibility assay and found that the hst1Δ mutant MIC50 was 6.3 times higher than BG14, consistent with the genetic data (Table 1 and Fig. 1A). Furthermore, given that S.c.Hst1 and C.g.Hst1 are highly similar (76%), we asked if S. cerevisiae mutant strains, Schst1Δ or Scsir2Δ, could also show reduced susceptibility to fluconazole. We found that S. cerevisiae parental strain (BY4742), Schst1Δ, and Scsir2Δ did not alter the fluconazole sensitivity to the extent caused by the hst1Δ mutant in C. glabrata (Fig. 1B). In summary, these results suggest that Hst1 acts as a negative regulator of genes that decrease susceptibility to fluconazole and that despite the high similarity between Sir2 and Hst1 (72%), these proteins have non-overlapping functions (Ma et al., 2009).
Table 1. Fluconazole susceptibility of C. glabrata strains
PDR1 and CDR1 are necessary for fluconazole decreased susceptibility in the absence of HST1
The overexpression of ABC efflux pumps represents the major mechanism responsible for high-level azole resistance in C. glabrata isolates (Bennett et al., 2004; Brun et al., 2004; Borst et al., 2005; Cannon et al., 2009; Ferrari et al., 2009). To date there is one report suggesting that genes of the pleiotropic drug resistance (PDR) could be negatively regulated by sirtuins (Ma et al., 2009). We decided to analyse the sensitivity to fluconazole when the transcriptional factor (PDR1) and the efflux pumps (CDR1 or CDR2) were deleted in the hst1Δ mutant background and in the presence of NAM. Consistent with previous reports (Sanglard et al., 1999; Vermitsky and Edlind, 2004), the pdr1Δ and cdr1Δ mutants were hypersensitive to fluconazole (Fig. 2A and Table 1, compare BG14, pdr1Δ and cdr1Δ). However, deletion of CDR2 (also known as PDH1; Izumikawa et al., 2003) did not alter the fluconazole sensitivity in the presence or absence of HST1 (Fig. 2A, compare BG14 and cdr2Δ and hst1Δ with cdr2Δ hst1Δ), possibly because Cdr1 is compensating for the absence of Cdr2 (Izumikawa et al., 2003). The decreased susceptibility to fluconazole of the hst1Δ strain was abolished in the absence of PDR1 or CDR1 (Fig. 2A, compare hst1Δ with pdr1Δ hst1Δ and cdr1Δ hst1Δ and Table 1). These results indicate that both genes are required to mediate azole resistance when Hst1 is absent. Notably, even when the pdr1Δ hst1Δ and cdr1Δ hst1Δ double mutants were more sensitive to fluconazole than the hst1Δ mutant strain, their sensitivity was not the same as the exhibited by the single mutant pdr1Δ or cdr1Δ (Fig. 2A, compare pdr1Δ with pdr1Δ hst1Δ and cdr1Δ with cdr1Δ hst1Δ), suggesting that CDR1 could be derepressed in pdr1Δ hst1Δ and PDR1 could be derepressed in cdr1Δ hst1Δ inducing the expression of other drug transporter encoding gene like CDR2; thus providing the additional reduced sensitivity. Quantification of the MIC50 of the single and double mutants indicated that the difference between these mutants is less evident than the genetic data (compare MIC50 of hst1Δ, pdr1Δ, and cdr1Δ, with hst1Δ pdr1∆ and hst1Δ cdr1Δ from Table 1 and Fig. 2A). The MIC50 value for the set of mutants shown in Table 1 was highly reproducible in YPD medium, however we could not confirm these phenotypes using RPMI-1640 medium as described in the NCCLS M-27 A2 method. Furthermore, the analysis of the susceptibility to fluconazole of the hst1Δ, pdr1Δ, cdr1Δ and the double mutants in the presence of NAM, were consistent with NAM inhibiting Hst1 activity. The presence of NAM mimics the absence of Hst1 (Fig. 2B, compare pdr1Δ and pdr1Δ hst1Δ with pdr1Δ hst1Δ -NAM in Fig. 2A and cdr1Δ and cdr1Δ hst1Δ with cdr1Δ hst1Δ -NAM in Fig. 2A). These data suggest that the decreased susceptibility due to the absence of HST1 is mainly mediated by CDR1 and PDR1.
The genetic evidence suggested that Hst1 could be a negative regulator of the expression of PDR1 and CDR1. In order to show, whether Hst1 is having a direct effect on the transcript levels of PDR1 and CDR1, we analyzed the expression of these genes by qPCR in both the parental strain and the hst1Δ mutant, treated with or without NAM. qPCR analysis showed that PDR1 and specially CDR1 transcript levels were higher in the hst1Δ mutant than in the parental strain (threefold and 16-fold increase, respectively, Fig. 3). Moreover, the expression of PDR1 and CDR1 also increased in the parental strain treated with NAM, consistent with the genetic data observed in Fig. 1A. These data indicate that Hst1 has a direct effect on the transcription of PDR1 and CDR1, acting as a repressor of their expression.
The absence of HST1 increases resistance to hydrogen peroxide and is dependent on MSN4 and CTA1
Several reports have suggested that a fast and efficient OSR may be important to support the survival of C. glabrata within the phagocytic cells (Cuellar-Cruz et al., 2008; Roetzer et al., 2010; Saijo et al., 2010; Seider et al., 2011). The regulators of the antioxidant response are beginning to be studied in this yeast and it has been shown that C. glabrata modifies substantially its epigenome during the phagocytosis (Rai et al., 2012).
Based on these observations, we decided to evaluate the involvement of Hst1 in the regulation of the OSR. We treated the parental strain BG14 with H2O2 as described in Experimental procedures. We found that hst1Δ mutant strain led to increased resistance to H2O2. The parental strain showed reduced growth at 50 mM of H2O2 while the hst1Δ strain was resistant up to 100 mM of H2O2 (Fig. 4A). This suggested that Hst1 could participate in the repression of genes of the OSR implicated in H2O2 detoxification. To assess this possibility, we constructed a double mutant strain lacking HST1 and the gene encoding the single catalase (CTA1), which is essential for H2O2 resistance in vitro (Cuellar-Cruz et al., 2008). We compared the sensitivity to H2O2 of the cta1Δ hst1Δ double mutant with the parental and the cta1Δ single mutant strain. The cta1Δ hst1Δ double mutant was slightly less sensitive to the oxidant as the cta1Δ mutant (Fig. 4A, compare cta1Δ and cta1Δ hst1Δ at 4 mM H2O2). Although CTA1 may be induced by several regulators (Roetzer et al., 2010), we decided to evaluate the involvement of the transcription factor MSN4 because it was overexpressed to a greater extent in a hst1Δ mutant strain (Ma et al., 2009). We constructed the msn4Δ hst1Δ double mutant strain and evaluated its resistance to H2O2. The msn4Δ mutant strain had the same sensitivity to H2O2 as the parental strain (Fig. 4A, compare msn4Δ with BG14). However, the msn4Δ mutation in the hst1Δ background abolished the H2O2 resistant phenotype. This result indicates that the oxidative stress resistance induced by the absence of HST1 is predominantly due to the presence of the catalase and might be regulated by Msn4. Furthermore, we also found that in S. cerevisiae, the response to oxidative stress is unchanged in the absence of ScHST1 or ScSIR2 (Fig. 4B). Along with the results described above, CgHst1 is regulating a different set of genes than those controlled by the orthologue ScHst1.
The genetic evidence suggests that the H2O2 resistance of the hst1Δ mutant could be due to a direct derepression of the catalase, and also that the induction of CTA1 could be mediated through the derepression of MSN4. To analyse these possibilities, we analysed the transcript levels of CTA1 and MSN4 in the parental and the hst1Δ mutant strain. Our qPCR analysis revealed a 3.5-fold increase in the expression of CTA1 in the absence of Hst1, and was only a subtle increase in CTA1 transcript levels in the presence of NAM (Fig. 5A). The expression of MSN4 increased fivefold in the presence of NAM, but only a twofold when Hst1 was absent (Fig. 5B). Given that these results suggest that the increase in resistance of the hst1Δ strain to H2O2 is due to an increase in catalase activity, we decided to quantify the activity of this enzyme (see Experimental procedures). Consistent with our results shown above, the catalase activity was 3.5-fold higher in the hst1Δ mutant than in the parental strain (Fig. 6). In contrast, enzymatic activity decreased in the hst1Δ msn4Δ double mutant strain, although not at the level exhibited by the parental or the msn4Δ mutant strains (Fig. 6). These results indicate that in the absence of HST1, C. glabrata is more resistant to H2O2 through the increase of the expression of CTA1 that requires the transcription factor Msn4.
Cysteine residues 344 and 347 are essential for Hst1 activity
Several amino acids have been identified as important for the structure and enzymatic activity of sirtuins in other organisms. Similar to other sirtuins, CgHst1 contains a Cys-X-X-Cys-(X)15–20-Cys-X-X-Cys conserved motif that co-ordinates a Zn2+ ion (Sherman et al., 1999). Interestingly, this same sequence motif (Cys-Pro-Tyr-Cys) is also located in the active site of glutaredoxins, which is a target for oxidation (Foloppe and Nilsson, 2004). We envisioned the possibility that Hst1 could be oxidized and inactivated in the presence of oxidative stress. To determine whether this sequence motif could be important for the activity of Hst1 in the presence of oxidative stress, we generated a mutant version of HST1 in which codons for cysteines 344 and 347 were substituted by alanines (see Experimental procedures). The substitution of this pair of amino acids was sufficient to eliminate Hst1 function, since the strain carrying the mutant allele (hst1 C344A C347A) was resistant to both oxidative stress and fluconazole at the same extent as the hst1Δ strain (Fig. 7 and Table 1). This result showed that the conserved motif Cys-X-X-Cys-(X)15–20-Cys-X-X-Cys is essential in CgHst1. Furthermore, to date there are no reports demonstrating inactivation of sirtuins by oxidation of specific residues, but this C-P-Y-C motif could be a potential target for oxidation.
Hst1 interacts with Sum1 to repress pleiotropic drug resistance genes and the oxidative stress response
In S. cerevisiae, Hst1 is tethered to the DNA-binding protein Sum1 through interactions with Rfm1 to form a complex that represses genes in a promoter-specific manner (Xie et al., 1999; McCord et al., 2003). In the C. glabrata genome, CAGL0J10956g correspond to CgSUM1 and CAGL0L11022g to CgRFM1 and interestingly, the amino acid sequence analysis revealed that CgRfm1 is 55% similar to ScRfm1 whereas CgSum1 is only a 23% similar to ScSum1. Even though we have shown that the Hst1 regulation of the fluconazole resistance genes and the regulation of OSR is specific to C. glabrata, we asked, if the Sum1-Rfm1-Hst1 complex could be present in this yeast. To determine genetically whether Sum1 and Rfm1 interact with Hst1, we generated single mutant strains in each gene and evaluated their susceptibility to fluconazole and H2O2. We found that the sum1Δ and rfm1Δ mutant strains showed the same decrease in susceptibility to fluconazole and H2O2 as the hst1Δ mutant strain (Fig. 8A and B). Moreover, the analysis of the double mutants, hst1Δ sum1Δ and hst1Δ rfm1Δ, showed no increase in the reduced susceptibility to fluconazole or H2O2 when compared with the single mutants, suggesting that these genes are components of the same pathway (Fig. 8A and B). As control, reconstitution of both the HST1 and SUM1 in their corresponding loci in the hst1Δ and sum1Δ mutant strains, reverted the fluconazole phenotype (data not shown). Finally and consistent with the results shown above for S. cerevisiae, deletion of ScHST1, ScSUM1 and ScRFM1 did not alter the susceptibility to fluconazole or H2O2 in this yeast (Figs 1B and 4B).
Our genetic analysis points toward an interaction between Hst1, Sum1 and Rfm1 in C. glabrata. In order to demonstrate this association, we performed a co-immunoprecipitation experiment using cMyc-tagged Hst1 and Flag-tagged Sum1 proteins. First, to determine if the tagged proteins were functional, we analysed the sensitivity to fluconazole of the HST1::cMyc SUM1::FLAG strain (CGM982). We found that the strain carrying the epitope-tagged alleles had the same fluconazole sensitivity phenotype as the parental strain, indicating that both proteins are functional (Fig. S2). We then performed the Co-IP using strain CGM982 (HST1::cMyc SUM1::FLAG) as described in the legend to Fig. 9 and in Experimental procedures. We found that Sum1-Flag co-immunoprecipitated with Hst1-Myc (Fig. 9, lane 4) and that this interaction is largely dependent on Rfm1, since the co-immunoprecipitation decreased when the RFM1 gene was deleted in the background of the double–tagged strain CGM1294 (Fig. 9, lane 5) and the majority of the Sum1-Flag remained in the supernatant. Taken together, our findings support a model in which Sum1, Rfm1 and Hst1 interact to form a complex that represses genes of the MDR and the OSR in C. glabrata.
Sirtuins are NAD+-dependent deacetylases highly conserved from bacteria to humans (Greiss and Gartner, 2009). In S. cerevisiae, sirtuins participate in gene silencing, DNA repair, replication, recombination, cell cycle control and longevity (Finkel et al., 2009; Vaquero, 2009). In the pathogenic yeast C. glabrata, sirtuins are involved in the transcriptional repression of genes encoding cell wall proteins which mediate adhesion and also genes encoding transporters of niacin, both processes are important for colonization of the host (Domergue et al., 2005; Ma et al., 2009). However, given the auxotrophy of C. glabrata for NAD+, the activity of sirtuins is completely dependent on the environmental availability of NAD+ precursors (niacin) (Ma et al., 2009). In host niches that are limited in niacin (urinary tract), sirtuins are inactivated and C. glabrata increases its adherence and niacin uptake. In this work, we decided to evaluate if inactivation of sirtuins modifies the capacity of this yeast to adapt to different stress conditions.
Inhibition of Hst1 decreases the susceptibility to fluconazole in C. glabrata
Here, we showed that C. glabrata decreases its susceptibility to fluconazole through the inhibition of Hst1 in the presence of NAM (Fig. 1). Based on our data, we propose that this phenotype is mediated primarily by the ABC transporter CDR1 and the transcription factor PDR1 (Figs 2 and 3; Table 1). However it is possible that other transporters could be derepressed as well, although with less affinity to fluconazole (Fig. 2). The main mechanism involved in the acquisition of resistance to azoles in C. glabrata, is the overexpression of genes belonging to the family of ABC transporters [Reviewed in (Cannon et al., 2009)]. Interestingly, to date almost all reported mechanisms involving overexpression of those transporters engage genetic modifications, such as the acquisition of gain of function mutations in Pdr1(Ferrari et al., 2009), loss of mitochondrial function (Kaur et al., 2004; Ferrari et al., 2011) and increased gene dosage of transporters by chromosomal rearrangements (Polakova et al., 2009). Recently it was demonstrated that some genes that provide azole resistance in C. glabrata are under the negative regulation of Stb5, a zinc cluster transcription factor (Noble et al., 2013). Here, we propose, based on our genetic and biochemical analysis, that Hst1 represses the transcription of PDR1 and CDR1. This hypothesis is further supported by the microarray analysis of the C. glabrata strain lacking Hst1 (Ma et al., 2009), where the absence of HST1 allows the induction of CDR1, PDR1 and CDR2 (PDH1).
Elimination of the Hst1-mediated silencing increases the oxidative stress resistance of C. glabrata
We found that C. glabrata increased the transcript levels of CTA1, its catalase activity and its resistance to H2O2 in the absence of HST1 (Figs 4, 5 and 6). Thus, it is probable that CTA1 is subject to negative control mediated by Hst1. The purpose of this negative regulation on the catalase is still unclear; although it has been shown that an increase in catalase activity is associated with premature aging of S. cerevisiae (Mesquita et al., 2010). This pro-aging activity has not been found in C. glabrata but silencing of CTA1 could contribute to maintain low level of transcript and limit the deleterious effect of this enzyme on viability. In addition, our results also suggest that the resistance observed in the absence of HST1 requires the transcription factor MSN4 (Figs 4 and 6). While it was true that the increase in the transcript levels of MSN4 were subtle, is likely to be sufficient to activate transcription of CTA1 (Fig. 5). Nevertheless, this finding apparently contrasts with previous observations that proposed a lack of participation by Msn2/4 in the control of CTA1 (Roetzer et al., 2011) although we detect the role of Msn4 only in the absence of Hst1. Thus, the resistance to oxidative stress observed in the hst1Δ cells may be due to at least two not mutually exclusive possibilities: 1) direct derepression of CTA1 or 2) derepression of one or several activators of CTA1 (such as MSN4) that are under the control of Hst1. Transcriptional regulation of CTA1 is a fine-tuned process that is subject to a complex interaction of factors that influence its activation and repression.
Sum1, Rfm1 and Hst1 repress genes of stress resistance in C. glabrata but not in S. cerevisiae
The precise mechanism by which Hst1 regulates the genes of the MDR and OSR is still not completely elucidated. However, here we showed that Hst1 could be repressing the transcription of genes that participate in both processes (Figs 3 and 5). Moreover, the putative transcriptional factor Sum1 and the connector protein Rfm1 were also required for the activity of Hst1 (Fig. 8) and the physical interaction of Hst1 with Sum1 was dependent on Rfm1 (Fig. 9). These findings are consistent with the results reported for S. cerevisiae (Xie et al., 1999; McCord et al., 2003) and suggest that the complex is conserved in both organisms. In S. cerevisiae, Hst1 and Sum1 control the NAD+ biosynthetic genes (Bedalov et al., 2003) and Hst1 has retained the role of regulating niacin uptake in C. glabrata (Ma et al., 2009). Surprisingly, deletion of any of the genes encoding the proteins that comprise the Hst1-Rfm1-Sum1 complex decreased susceptibility to fluconazole and H2O2 in C. glabrata but not in S. cerevisiae (Figs 1 and 4). These similar deacetylases (74% similar) control different genes in phylogenetically related organisms. In agreement with this observation, the transcriptional profiles of S. cerevisiae and C. glabrata strains carrying a mutation in HST1, show that only 19 genes are regulated in common by this sirtuin in these two yeasts [Comparing the upregulated genes found in the microarrays reported by Ma and Hirao (Hirao et al., 2003; Ma et al., 2009)]. It is possible that the reason for this differential regulation might lie in Sum1. At the protein level, CgSum1 is only 23% similar to ScSum1. BLAST analysis shows no putative conserved motifs over the entire length of Sum1 proteins, but the N-domain appears to be the most conserved region in C. glabrata and other phylogenetically related yeasts (data not shown). Despite the divergence that CgSum1 has undergone, our results suggest that it may conserve important amino acid residues needed for its interaction with CgRfm1 and probably also for DNA binding. Based in our data we propose a model in which Sum1 interacts with Rfm1 and directs Hst1 to the promoters of the MDR and OSR genes to silence them (Fig. 10).
Hst1 as a regulator of genes associated with survival within the host
Our data also confirm that despite the high similarity between Sir2 and Hst1 (74%), only the absence of Hst1-mediated silencing decreases the susceptibility of C. glabrata to fluconazole and oxidative stress. In addition, deletion of HST1 also increases the in vitro adherence of C. glabrata to Lec2 epithelial cells (B.P. Cormack and I. Castano, pers. comm.) and the transcription of EPA6 (Domergue et al., 2005), suggesting that Hst1 also participates in the repression of some epithelial adhesins. Therefore, it appears that Hst1 represses genes that could be advantageous to this pathogen (Fig. 10), thus Hst1 could be inactivated during infection. C. glabrata colonizes the genitourinary tract where the levels of niacin are limited so it is feasible that Hst1 loses its activity and allow derepression of its target genes under this condition. Consistent with this, it has been suggested that MSN4 and CTA1 are upregulated under limitation of niacin and in human urine (Ma et al., 2009).
According to our model, inactivation of Hst1 could be an advantage for C. glabrata because it would allow colonization of the host through the derepression of specific adhesins and also would increase resistance to xenobiotics and oxidative stress (Fig. 10). This work uncovers a novel level of regulation in the MDR and OSR circuits of C. glabrata. We think that this epigenetic regulation of resistance could be advantageous for C. glabrata because, either it might allow the stochastic appearance of resistant cells within a genetically identical population without permanent genetic alterations that may compromise the fitness of the yeast or the Hst1-Rfm1-Sum1 complex could be a target of different environmental signals. It remains to be evaluated whether these mechanisms are relevant to the acquisition of resistance to fluconazole during therapy of the patients and to endure the respiratory burst during phagocytosis. It will also be worth exploring whether maintaining the activity of Hst1 could be beneficial to the host, as this condition would keep silent the genes that mediate adhesion and resistance of C. glabrata.
Strains and growth conditions
Escherichia coli strain DH10 (Gibco BRL) was used for plasmid transformation, isolation and storage in 10% glycerol. All mutant C. glabrata strains (Table 2) were generated in the BG14 strain background and here referred to as parental strain (Cormack and Falkow, 1999). The S. cerevisiae mutant strains were all derivatives from BY4742 (Table 3). All yeast strains were stored in 15% glycerol stocks at −80°C. Yeast cultures were incubated at 30°C in YPD [1% yeast extract, 2% peptone, 2% glucose and supplemented with 25 mg ml−1 of uracil (Sigma Aldrich®)] or in synthetic complete medium (SC) (yeast nitrogen base at 1.7 g l−1 and (NH4)2SO4 at 5 g l−1, supplemented with 0.6% Casamino Acids and 2% glucose). For solid media, 2% agar was added.
Plasmids used in this study are described in Table S1. All plasmids were purified from E. coli using the Qiagen® Plasmid Kit QIAprep. All gene fragments for plasmid constructs and for gene disruption were generated by PCR using the oligonucleotides detailed in Table S2. Fusion PCR for gene disruption was performed as previously described (Baudin et al., 1993) and all PCR products were purified before transformation using Qiagen® Gel Extraction Kit. C. glabrata was transformed using the lithium acetate protocol and the knockout mutations and the epitope-tagging were constructed by two-step recombination; these methods had been described previously (Castano et al., 2003; De Las Penas et al., 2003; Cuellar-Cruz et al., 2008). To construct the null mutant strains, the open reading frame (ORF) of each target gene was replaced with a hygromycin resistance cassette or with the nourseothricin resistance cassette. The plasmids used for epitope tagging contain a C-terminal segment of the gene without stop codon (HST1 or SUM1) placed in phase with the epitope sequence (cMyc or FLAG). This translational fusion is followed by the 3′ UTR of the catalase gene (3′ UTRCTA1) and a hygromycin resistance cassette for selection of the transformants. The 3′ UTRCTA1 and the hygromycin cassette are flanked by two FRT direct repeats. Downstream the hygromycin cassette, the constructs contain a fragment of the 3′ UTR of the gene. The mutant or epitope tagged strains were selected on YPD plates containing 440 μg ml−1 hygromycin (A. G. Scientific®) or 100 μg ml−1 nourseothricin (Werner BioAgents®). The correct replacement in the chromosome was confirmed by PCR. For each target gene, two independent transformants were obtained and tested. To place the HST1-cMyc and SUM1-FLAG with their native 3′ UTR, the epitope-tagged HygR strains were transformed with pMZ21 (Table S1), and transformants were selected on SC-Ura plates. pMZ21 is a replicative vector expressing ScFLP1 that encodes a recombinase that recognizes the FRT sites flanking the hygromycin marker. After Flp1 recognizes the FRT sites, the hph marker is deleted from the chromosome and the cognate 3′ UTR is placed immediately downstream of the tagged gene and leaves one copy of the FRT site (35 bp). Transformants were purified on SC-Ura plates. Single colonies were then grown on non-selective medium (YPD agar) and screened for HygS for the loss of the Hyg cassette and for Ura− for the loss of pMZ21. This protocol allows the construction of multiple mutants or multiple tagged strains.
For site directed mutagenesis, the coding region of HST1 was mutated using PCR so that Cys residues were replaced by Ala. Expand Long Template PCR System (Roche®) was used for amplification of the fragments. Sequences of the mutagenic oligonucleotides used for the generation of the hst1 C344A C347A mutant are listed in Table S2. The mutagenic oligonucleotides were designed to mutate the selected codons and simultaneously introduce a novel Nhe I restriction site, which facilitates distinction of mutant from wild-type fragment at subsequent cloning steps. To confirm the introduction of the mutation, plasmid pOZ50 was sequenced. To replace HST1 with the mutant allele (C344A C347A), WT-strain was transformed with linearized pOZ86 using the pop-in/pop-out method. Segregants were selected on SC-Ura plates. To promote recombination, colonies were cultured in YPD broth and plated on YPD agar plates. Loss of URA3 was selected by streaking colonies on 5-FOA plates. Finally identification of the mutants was achieved using PCR, restriction analysis and sequencing.
Fluconazole susceptibility assays
To test for susceptibility to fluconazole (Pfizer®), C. glabrata cells were grown for 48 h at 30°C in YPD broth, diluted to OD600 = 0.5 in distilled water, and 10-fold serial dilutions were spotted onto YPD agar plates containing different fluconazole concentrations (Sanglard et al., 1999). To assess fluconazole susceptibility under inhibition of sirtuins, 10 mM of NAM (Sigma Aldrich®) was added to the plates containing fluconazole. Spots were made using a VP 407AH 48-pin multi-blot replicator (V&P Scientific Inc.). Plates were incubated at 30°C for 36 h or 48 h. Experiments were repeated at least three times.
To quantify the sensitivity to fluconazole, stationary phase cultures of the indicated strains were diluted to obtain a cellular suspension in YPD. One hundred and fifty microlitres of this suspension (104 cells ml−1) was transferred to multi-wells plates (Oy Growth Curves Ab Ltd) where each well contained 150 μl of broth with different concentrations of fluconazole (0.0078, 0.0156, 0.0312, 0.0625, 0.125, 0.250, 0.5, 1, 2, 4, 8, 16, 32, 64, 72, 80, 96, 112, 128, 160, 192, 224 or 256 μg ml−1). Growth was then continuously monitored in a Bioscreen C system (Oy Growth Curves Ab Ltd) at 600 nm, for each 15 min at 35°C during 24 h. Sensitivities were determined as mean 50% Minimal Inhibitory Concentrations (MIC50), calculated on the basis of the dose-response relationship in the GraphPad Prism software (GraphPad Software Inc., San Diego, CA, USA). The CLSI microdilution method M27-A2 was also used for susceptibility testing but RPMI-1640 broth was supplemented with 25 mg ml−1 of uracil.
H2O2 susceptibility assay
Sensitivity to hydrogen peroxide (H2O2) of logarithmic phase cells was determined as described previously (Cuellar-Cruz et al., 2009). A 35% wt H2O2 solution (Sigma Aldrich®) was used for the assay. Cultures were grown for 48 h at 30°C in YPD broth, diluted into fresh YPD medium in such way that all the strains went through seven duplications to reach 0.5 OD600. After this period, the cultures were divided, exposed to different H2O2 concentrations and incubated with shaking at 30°C for three hours. After the treatment, the H2O2 was removed by centrifugation and the cells were suspended in 1 ml of distilled water. The cultures were adjusted to an OD600 = 0.5, serially diluted and spotted on YPD plates as described above. The plates were incubated at 30°C for 36 h or 48 h. The experiments were performed at least three times.
Catalase activity assay
Strains were grown at 30°C for seven duplications in YPD broth until cultures reached an OD600 = 0.5. Cells were collected by centrifugation, washed twice with distilled water and suspended in lysis buffer [pottasium phosphate (pH 7.0) supplemented with 1× general protease inhibitors [Sigma FAST®]). A volume of zirconia beads (BioSpec®) was added and cells were disrupted by vortexing at 4°C for 1 min and placed on ice for another minute (repeated 20 times). The lysate was centrifuged at 25 000 g for 30 min at 4°C to remove cell debris and zirconia beads. The supernatant was used for measure of catalase activity and its protein content was determined by Bradford assay (Fermentas®). Bovine serum albumin (Sigma®) was used as standard. Catalase activity was determined by the spectrophotometric method that measures the breakdown of H2O2 by catalase (Aebi, 1984) and the assays were performed three times. The activity was normalized to total content of protein from the lysate and expressed as units per mg of protein. One unit is defined as the amount of catalase required for degradation of 1.0 μmol H2O2 per minute.
Quantitative PCR (qPCR)
RNA was extracted from log phase cells (grown in YPD or in YPD broth containing 10 mM NAM) using TRIzol reagent (Invitrogen) according to the manufacturer's instructions and treated with DNase I (Invitrogen) to ensure removal of contaminating genomic DNA. Synthesis of cDNA was carried out using SuperScript® II reverse transcriptase and the reverse oligonucleotides for each gene (Table S2) and the reaction was carried out at 42°C for 50 min. Quantitative PCR (qPCR) was performed using the Fast SYBR® Green Master Mix (Invitrogen) in the ABI 7500 Fast Real-Time PCR System (Applied Biosystems). The reverse and forward oligonucleotides used for the qRT-PCR were designed using the primer express 3.0 software (Table S2, ACT1 #41–42, CTA1 #43–44, CDR1 #45–46, MSN4 #47–48 and PDR1 #49–50). ACT1 was used as an internal control for normalization and the threshold cycle (2−ΔΔCt) method was used to calculate the differences in gene expression. qPCR experiments were performed in technical triplicates and from three independent RNA extractions.
The CoIP protocol was performed as described (McCord et al., 2003). Strains were grown at 30°C for seven duplications in YPD broth until cultures reached an OD600 = 1.0. Cells were collected by centrifugation and suspended in 300 μl of lysis buffer (45 mM HEPES-KOH [Promega®, pH 7.5], 400 mM potassium acetate, 1 mM EDTA, 0.5% Nonidet P-40 substitute [Fluka Biochemica®], 1 mM DTT, 10% glycerol, 1 mM PMSF, 1 × general protease inhibitors [Sigma FAST®] and 1 × Complete protease inhibitors [ROCHE®]). A volume of 300 μl of zirconia beads (BioSpec®) was added and cells were lysed as described for in the catalase activity protocol. The lysate was centrifuged at 25 000 g for 30 min at 4°C. The supernatant of each sample was transferred to a new tube and protein content was determined by Bradford assay (Fermentas®). For each sample, 100 μg of protein were incubated at 4°C for 1 h with 30 μl of sepharose beads containing cross-linked protein G (Sigma®). The samples were centrifuged at 25 000 g for 5 min at 4°C and the clarified lysate was transferred to a clean tube. For immunoprecipitation, the lysate was combined with 50 μl of anti-cMyc or anti-FLAG agarose (Sigma®) and incubated at 4°C for two hours with constant inversion. The supernatant was recovered and IPs were collected by centrifugation, washed three times with Lysis buffer, suspended in 40 μl of 2× sodium dodecyl sulphate (SDS) loading buffer and heated at 95°C for 5 min. The input (20 μg of protein), supernatant (one twentieth of the immunoprecipitated extract) and the IPs were loaded onto a 10% SDS-polyacrylamide gel. After electrophoresis, the proteins were blotted onto PVDF membranes (BIO-RAD®) and probed with either mouse anti-cMyc (Millipore®) or mouse anti-FLAG (Sigma®) for two hours at room temperature. After washing, the membranes were probed with a goat anti-mouse horseradish peroxidase-conjugated secondary antibody (Amersham®). Signal detection was achieved using the ECL chemiluminescence reagents (Amersham®) and X-OMAT (Kodak®) films.
We thank Jasper Rine for kindly providing S. cerevisiae strains (University of California, Berkeley, CA, USA). We are grateful to Sergio Casas and Lina Riego (IPICYT, San Luis Potosí, SLP, Mexico) for providing reagents. We thank Candy Y. Ramírez, Omar Arroyo and Marcela Briones for technical support, Jorge Folch (UAEM) and members of the A.D.L.P. laboratory for their suggestions and careful reading of the manuscript. This work was funded by CONACYT fellowships to E.O.Z. (233455), G.G.S. (230938), M.G.G.E. (48880), I.C.V. (224300), J.J.C. (48549) and by CONACYT Grant No. CB-2010-01-153929 and UC Mexus Grant No. CN-07-53 to A.D.L.P.