Browse Subject Areas

Click through the PLOS taxonomy to find articles in your field.

For more information about PLOS Subject Areas, click here.

  • Loading metrics

Role of Ess1 in Growth, Morphogenetic Switching, and RNA Polymerase II Transcription in Candida albicans

  • Dhanushki Samaranayake,

    Affiliations Department of Biomedical Sciences, School of Public Health, State University of New York, Albany, New York, United States of America, Division of Genetics, Wadsworth Center, NY State Department of Health, Albany, New York, United States of America

  • David Atencio,

    Affiliation Department of Biochemistry and Molecular Biology, State University of New York, Upstate Medical University, Syracuse, New York, United States of America

  • Randall Morse,

    Affiliations Department of Biomedical Sciences, School of Public Health, State University of New York, Albany, New York, United States of America, Division of Genetics, Wadsworth Center, NY State Department of Health, Albany, New York, United States of America

  • Joseph T. Wade,

    Affiliations Department of Biomedical Sciences, School of Public Health, State University of New York, Albany, New York, United States of America, Division of Genetics, Wadsworth Center, NY State Department of Health, Albany, New York, United States of America

  • Vishnu Chaturvedi,

    Affiliations Department of Biomedical Sciences, School of Public Health, State University of New York, Albany, New York, United States of America, Mycology Laboratory, Wadsworth Center, NY State Department of Health, Albany, New York, United States of America

  • Steven D. Hanes

    Affiliations Department of Biomedical Sciences, School of Public Health, State University of New York, Albany, New York, United States of America, Division of Infectious Disease, Wadsworth Center, NY State Department of Health, Albany, New York, United States of America, Department of Biochemistry and Molecular Biology, State University of New York, Upstate Medical University, Syracuse, New York, United States of America

Role of Ess1 in Growth, Morphogenetic Switching, and RNA Polymerase II Transcription in Candida albicans

  • Dhanushki Samaranayake, 
  • David Atencio, 
  • Randall Morse, 
  • Joseph T. Wade, 
  • Vishnu Chaturvedi, 
  • Steven D. Hanes


Candida albicans is a fungal pathogen that causes potentially fatal infections among immune-compromised individuals. The emergence of drug resistant C. albicans strains makes it important to identify new antifungal drug targets. Among potential targets are enzymes known as peptidyl-prolyl cis/trans isomerases (PPIases) that catalyze isomerization of peptide bonds preceding proline. We are investigating a PPIase called Ess1, which is conserved in all major human pathogenic fungi. Previously, we reported that C. albicans Ess1 is essential for growth and morphogenetic switching. In the present study, we re-evaluated these findings using more rigorous genetic analyses, including the use of additional CaESS1 mutant alleles, distinct marker genes, and the engineering of suitably-matched isogenic control strains. The results confirm that CaEss1 is essential for growth in C. albicans, but show that reduction of CaESS1 gene dosage by half (δ/+) does not interfere with morphogenetic switching. However, further reduction of CaEss1 levels using a conditional allele does reduce morphogenetic switching. We also examine the role of the linker α-helix that distinguishes C. albicans Ess1 from the human Pin1 enzyme, and present results of a genome-wide transcriptome analysis. The latter analysis indicates that CaEss1 has a conserved role in regulation of RNA polymerase II function, and is required for efficient termination of small nucleolar RNAs and repression of cryptic transcription in C. albicans.


Candida albicans causes life-threatening fungal infections in hospitalized patients [1][3]. C. albicans is a commensal organism found on the human mucosal surface and is generally harmless in healthy individuals [4], [5]. However, C. albicans can cause systemic and sometimes fatal infections in immune-compromised individuals [6][8]. Life-saving therapies that require suppression of the immune system, e.g. organ transplantation and cancer chemotherapy, increase the risk for invasive candidiasis [9][12]. Premature infants, HIV-infected individuals, and individuals receiving prolonged intensive care treatment or antibiotic treatment are also vulnerable [13]-[16]. While potent antifungal drugs are available, the emergence of drug-resistant strains, especially against the widely used azole drugs is a growing problem [3], [17][21].

One strategy to overcome drug resistance while producing synergistic effects is the use of combination therapies that target distinct intracelluar pathways [22][27]. Toward this end, a number of different pathways are being investigated including those containing enzymes known as peptidyl prolyl cis-trans isomerases [28], [29]. PPIases catalyze the isomerization of the peptide bond preceding prolines within protein substrates [30], [31]. Three major families of PPIase have been described; cyclophilins, FK506-binding proteins (FKBPs) and parvulins (reviewed in [32]. All three families are conserved from yeast to humans. Inhibitors such as cyclosporin, which targets the cyclophilins, and FK506 or rapamycin, which target FKBPs, all show potent antifungal activity [33][36]. However, these drugs are also immunosuppressive via pathways that inhibit T-cell activation, an activity that could make individuals more vulnerable to fungal infections [37], [38].

An alternative PPIase target might be Ess1, a founding member of the parvulin class of PPIase, which is structurally distinct from the cyclophilins and FKBPs [39], [40] and whose human ortholog Pin1 is not known to be associated with the T-cell activation pathway. The first fungal Ess1 was discovered in Saccharomyces cerevisiae and shown to be essential [41]. Ess1 and Pin1 play critical roles in gene transcription by RNA polymerase II (pol II) [42][46]. Ess1 isomerizes peptide bonds within the carboxy-terminal domain (CTD) of Rpb1, the largest subunit of RNA pol II, and thereby controls binding and release of transcriptional co-factors [43], [47][49]. The X-ray structures of the C. albicans Ess1 protein and its human ortholog, Pin1 have been solved [40], [50], [51], and while the enzymes show overall similarity, there are key differences including a large solvent-exposed alpha-helix within a structured linker region that is present in the fungal enzyme but absent in the human enzyme. This helix has been hypothesized to play a role in fungal-specific functions, potentially by engaging in protein-protein interactions [50].

Homologs of Ess1 are found in all major pathogenic fungi that have been examined, including C. albicans [52], Candida glabrata (REFSEQ XP_445146), Cryptococcus neoformans [53], and Aspergillus nidulans (PinA) [54]. In C. albicans and A. nidulans Ess1 (PinA) is essential for growth [52], [54]. In Cryptococcus neoformans, Ess1 is not essential for growth, but is required for expression of virulence factors melanin and urease and for virulence in a mouse model [53].

In previous work, we isolated the CaESS1 gene from C. albicans and showed it to be essential for growth in this organism using a novel temperature-sensitive mutant strategy [52]. Surprisingly, we found a heterozygous mutant strain (Caess1δ/CaESS1) to be defective for filamentation in various inducing media leading us to conclude that CaESS1 gene dosage is important for morphogenetic switching [52]. However, several considerations led us to re-evaluate these findings. First, our CaESS1 dosage experiments used the URA-blaster method [55], which has largely been superseded to avoid variations in virulence phenotypes due to expression of URA3 from ectopic loci [56], [57]. Second, the importance of using reconstituted strains rather than parental strains as controls is now well-established [57],[58]. Finally, sequencing of the C. albicans genome [59] revealed a gene, APE2 located very close to CaESS1, whose promoter might have been disrupted in our previous strain constructions.

We therefore re-examined our previous findings using newer information and more rigorous methods. The results showed that C. albicans Ess1 is essential for growth, but that in contrast to our previous report, CaESS1 heterozygous mutants did not show a defect in morphogenetic switching, and instead the defect was traced to a host cell mutation(s). Potential effects of URA3 marker gene placement were also ruled out. We also report results of a structure-function analysis of the CaEss1 linker α-helix, and describe a conditional-lethal readthrough allele of CaESS1 that will be useful for further studies. Finally, results of a transcriptome analysis using high-throughput RNA-sequencing suggests that C. albicans Ess1, like its counterparts in budding yeast and humans plays a role in regulating RNA polymerase II function.

Results and Discussion

Reduction of CaESS1 gene dosage does not affect filamentation phenotypes

In previous work, we constructed a C. albicans Caess1δ/CaESS1 heterozygous mutant strain in the CAI4 (ura) parent strain using the URA-Blaster method [55] (Figure 1A). This strain, CaGD1, was defective for filamentation in Lee's, serum-containing and Spider media when compared to the “wild-type” control SC5314 [52]. The CAI4 (ura) parent strain is a poorly-filamenting strain, presumably due to its uracil auxotrophy [60]. Instead, the parent strain of CAI4, clinical isolate SC5314, had been used as a control, because SC5314 is a uracil prototroph (ura+) and could therefore be compared to CaGD1 (ura+), even though SC5314 has two copies of URA3 and CaGD1 has only one copy.

Figure 1. Schematic representation of the strains used in this study.

Strains AH are made in the CAI4(*) background using URA3 as a selectable marker. Strains IO are made in SN87 background with HIS1 and LEU2 selectable markers. The CaESS1 coding region and promoter region (pCaEss1) are shown in green; the APE2 gene is shown in purple. APE2 gene has two exons and the wavy line represents the intronic region. CaESS1 and APE2 are 210 bp apart in the genome (grey). The URA3 gene (blue) with the two flanking hisG direct repeats from S. typhimurium (yellow) is part of the construct used in the URA-Blaster method to target genes and later recycle URA3. RPS1 is shown in brown, LEU2 in red, HIS1 in pink and (p) indicates promoter sequence. The sequence upstream of APE2 that is repeated is shown in a gray-white gradient. The CAI4(*) indicates that this strain appears to have acquired a mutation that affects filamentation as per this study. Figure is not drawn to scale.

Here, we generated a better control, isogenic to CAI4, but with one copy of the URA3 gene placed at the CaESS1 locus without disrupting it (Figure 1E). Surprisingly, this reconstituted control strain, R6 (ura+) (CaESS1/CaESS1), also did not filament on serum-containing medium or Spider medium even though both CaESS1 alleles were left intact (Figure 2E). Filamentation on Lee's medium was also defective (data not shown). It is possible that the integrated URA3 construct in R6 somehow lowered CaESS1 expression levels in cis, but this was ruled out using quantitative reverse transcription real time PCR (qRT-PCR) and Western analysis, which shows the expected CaESS1 RNA and protein levels in the heterozygotes (CaGD1) and wild-type control (R6) strains (Figure 3). Thus, placement of the URA3 gene at the CaESS1 locus or some other defect, but not CaESS1 dosage, was likely responsible for the filamentation defect.

Figure 2. Filamentation of strains used in the study.

Filamentation was tested on solid medium at 37°C. (Upper panel) Serum medium (4% FBS in 2% agar), (Lower panel) Spider Medium [80]. 2 µl of the 0.25 OD600 of fresh overnight cultures were spotted and grown for 4 days at 37°C before documentation. Small letters (A–O) within each panel refer to the constructs shown in Figure 1.

Figure 3. CaESS1 is expressed at reduced levels in heterozygous mutants as expected.

(A) Quantitative reverse transcription PCR (qRT-PCR) shows CaESS1 mRNA expression levels in the indicated strains. (B) Western blot analysis showing the expression of CaEss1 protein in the indicated strains. A total of 3 µg of protein was used per lane. The blot was probed using anti-CaEss1 polyclonal antibody at a 1:500 dilution. CaEss1 is ∼19 kD. Strains correspond to those shown in Figures 1 and 2: (ESS1/ESS1) URA3 at CaESS1 is R6; (ess1Δ/ESS1) URA3 at CaESS1 is CaGD1; (ESS1/ESS1) HIS1/LEU2 is CaDS-B5; (ess1Δ/ESS1) HIS1/LEU2 is CaDS-B5.5.

One possibility was that the URA3 gene placed at the CaESS1 locus interrupted the promoter region of the downstream gene, APE2. The APE2 gene encodes an aminopeptidase that is thought to be secreted through the cell wall [61]. The design of our URA3-based CaESS1 knockout and control constructs leaves only 15 bp of upstream sequence on one allele of APE2, leaving open the possibility that reduced expression of APE2 was responsible for the filamentation defect in both strains (Figure 1A, 1E). As described in a later section, neither URA3 nor APE2 expression was involved.

To independently confirm that CaESS1 gene dosage did not affect filamentation we took an approach that did not rely on URA3 markers or disrupt the APE2 promoter. We used a parent strain (SN87) in which HIS1 and LEU2 are used as selectable markers [62]. Extensive studies have shown that ectopic expression of these markers does not significantly affect the virulence functions [62]. Accordingly, we replaced one allele of CaESS1 (with HIS1) generating a heterozygous strain (the other allele was marked with LEU2) (Figure 1I). We also constructed a control strain in the same background, but leaving both CaESS1 alleles intact (Figure 1J). To avoid any expression defects in CaESS1 or the downstream APE2 gene, the constructs were designed to have a short direct-repeat sequence (163bp) flanking the markers such that the ESS1 gene would have a total of 200 bp downstream of the coding sequence for transcription termination, and the APE2 gene would have 173bp of promoter sequence left intact (Figure 1I, J; see also Materials and Methods). As before, qRT-PCR and Western analysis shows the expected reduction by about half of the expression of CaESS1 mRNA and protein in heterozygous mutants (CaDS-B5.5) vs. the isogenic wild-type control (CaDS-B5 (Figure 3).

The Caess1δ/CaESS1 heterozygous mutant strain and its CaESS1/CaESS1 isogenic control showed no significant differences in filamentation phenotypes on inducing media (Figure 2I, J). In addition, no difference between these strains was observed in either germ-tube formation assays or in drug susceptibility growth assays using a number of commercially available antifungal drugs (data not shown). In summary, we find that the reduction of CaESS1 gene dosage by half does not affect major virulence-related phenotypes of C. albicans in-vitro. We conclude that one copy of CaESS1 is sufficient for growth and morphogenetic switching consistent with the finding that Ess1 in S. cerevisiae is present in excess under standard growth conditions [47].

Neither URA3 nor APE2 levels are responsible for the filamentation defect in our CAI4 isolate

If CaESS1 gene dosage was not responsible for the filamentation defects in our CAI4-derived strains, then what was responsible? To determine if ectopic URA3 at the ESS1 locus caused the filamentation defects in CaGD1 (Caess1δ/CaESS1) [52] and R6 (CaESS1/CaESS1) strains (Figure 2A,E), we created two additional sets of strains. In one set, the URA3 gene in both the CaGD1 and R6 strains were removed and replaced into the native URA3 locus (Figure 1B,C & F,G). In the other set, the URA3 gene was removed and placed at the RPS1 locus (Figure 1B,D & F,H). The RPS1 locus is commonly used to express URA3 and other genes to avoid positional affects [56], [57], [62]. We then used qRT-PCR to compare the levels of expression of URA3 that resided at the CaESS1, native (URA3) or RPS1 loci. This was done for cells grown in selective (uracil-deficient) media at 30°C and in 10% serum medium (10% FBS in YPD) at 37°C (Figure 4A,B). URA3 expression was essentially the same regardless of where the gene resided, thus ruling out position effects on URA3 expression as causing filamentation defects in our CAI4-derived strains. Moreover, placement of URA3 at different locations had no effect on expression of CaESS1 mRNA or protein (Figure 3).

Figure 4. URA3 and APE2 mRNA expression levels are not significantly altered in heterozygous ESS1 mutants and controls.

qRT-PCR showing a quantitative measurement of URA3 and APE2 mRNA expression in the indicated strains. (A, C) Cells were grown at 30°C in complete synthetic medium (CSM), CSM minus uracil, or CSM minus histidine and leucine as appropriate for each strain. (B, D) Cells were grown at 37°C in serum-containing medium (10% FBS in YPD). Strains are as described in legend to Figure 3.

Note, however, that all of the above strains carry one copy of the APE2 gene with a truncated or hisG-disrupted promoter (Figure 1AF). To test whether reduced APE2 expression caused the filamentation defect, APE2 expression was measured in multiple strains (corresponding to Figure 1AJ) using qRT-PCR. Expression was measured in cells grown in selective media (30°C) and in serum-inducing medium (10% FBS in YPD, 37°C), and no significant differences in APE2 expression were detected (Figure 4C,D). These results suggest that the remaining copy of APE2 with an intact promoter (strains in Figure 1AF) compensated for the loss of upstream sequences in the second copy, or that the transcription is regulated from an internal promoter. In experiments done in the SN87 background using a strain with both APE2 promoter regions truncated (5′ape2/5′ape2) (Figure 1K) neither the expression of APE2 (data not shown) nor filamentation (Figure 2K) is affected. Results thus far indicate that APE2 expression is not correlated with the filamentation defect in CAI4-derived strains.

Finally, to characterize our parental CAI4 isolate, which we now concluded was suspect, we placed URA3 at the native locus or at the RPS1 locus and tested its ability to filament. Analogous CAI4-derived strains (e.g. CAI12, Figure 2R) made in other laboratories using similar approaches [56], [63], [64] were able to filament, but the strains derived from our CAI4 isolate were not (Figure 2P,Q). Thus, not all CAI4 isolates behave similarly under inducing conditions, and we suspect that the isolate we used previously had acquired a background mutation(s) that resulted in a loss in the ability to undergo morphogenetic switching. These results explain the difference between our current results and those reported in Devasahayam et al., [52] with respect to CaESS1 gene dosage and filamentation phenotypes. They may also explain the reduction in organ load in mice injected with SC5314 vs. CAI4-derivatives with mutations in CaESS1 [50].

CaESS1 is essential for growth in C. albicans

Previously, a novel temperature-sensitive (ts) approach was used to demonstrate that CaESS1 is essential for C. albicans viability [52]. In this approach one allele of CaESS1 was deleted and the second allele was replaced with a form of CaESS1 engineered to be conditional. Specifically, a substitution in a critical histidine residue in the C. albicans Ess1 active site (H171R) was generated based on a ts-mutation (ess1H164R) well characterized in S. cerevisiae [42]. Prior to its use in C. albicans, the CaESS1H171R allele was tested in complementation assays in S. cerevisiae and confirmed to be conditional; cells grew at 25°C and 30°C, but not at 37°C. When this allele was integrated into C. albicans CAI4, the resulting strain (Caess1δ/Caess1H171R strain) was ts-lethal: it grew at 30°C but not at 40°C, demonstrating that Ess1 is essential in C. albicans.

Here we sought to confirm, using this ts-strategy, that Ess1 is essential in another strain background, using a marker system other than URA3. Despite repeated attempts in the SN87 background, we were unable to generate a Caess1Δ/Caess1H171R strain, consistent with CaEss1 being essential and suggesting that in this strain background the Caess1H171R allele cannot support growth. However, this is a negative result, so we took another approach.

We noted that the analogous S. cerevisiae ess1H164R protein has<0.01% the catalytic activity of wild-type protein [47] and decided to generate an additional allele, again based on prior work in S. cerevisiae [42]. The allele we generated alters the termination codon (TAA to TGC), resulting in translational readthrough. In S. cerevisiae, analogous mutations resulted in longer, fusion proteins that rendered the corresponding strains ts (X. Wu and S. Hanes, unpublished). The C. albicans strain we generated here, CaDS-C (ess1δ/Caess1TGC; Figure 1L), which contains mutations in the stop codon (TAA to TGC) in CaEss1, encodes a protein that is larger than normal and is thermolabile at 42°C (Figure 5A). The level of mutant protein is only slightly reduced after two hours at 37°C (data not shown).

Figure 5. CaESS1 is essential for the growth of C. albicans.

(A) Western analysis of C. albicans whole cell extracts to detect expression of wild-type CaEss1 (TAA) and a larger protein product encoded by the Caess1TGC “readthrough” allele (TGC). The level of read-through protein is reduced significantly at 60 min and is nearly absent after 2 hrs at non-permissive temperature (42°C). A total of 7.5 µg of protein was used per lane. The blot was probed using anti-CaEss1 polyclonal antibody at a 1∶500 dilution. (B) Serial dilution (1∶3) of cells of the indicated genotype grown on solid medium (YPD) at different temperatures. The readthrough strain, Caess1Δ/Caess1TGC shows a clear temperature-sensitive phenotype at 42°C, but no growth defect at 37°C. (C) Filamentation on the indicated solid medium (4 days) (upper two rows), and germ tube formation in liquid Spider medium (2 hrs) (lower row). Upper two rows are reproduced from Figure 2 for comparison. In (B) and (C), CaDS-B5 (CaESS1/CaESS1) is used as the wild-type, and is an isogenic control for both the TAA and TGC strains.

We compared the growth at different temperatures of the termination-mutant strain (CaDS-C) to an isogenic, reconstituted control strain in which the TGC readthrough codon was changed back to a TAA termination codon (CaDS-FC; Caess1δ/CaESS1TAA). Both strains grew at lower temperatures (30°C and 37°C), but only the wild type (reconstituted) strain grew at 42°C (Figure 5B). The results show that the Caess1ts(TGC) allele is unable to support sustained growth at the restrictive temperature demonstrating that CaEss1 is essential for growth in the SN87 strain background. Thus, using a distinct conditional allele (Caess1TGC) in a different strain (SN87) with different markers (HIS1, LEU2), we confirm that CaEss1 is essential in C. albicans.

A large reduction in Ess1 levels reduces filamentation

In the course of characterizing the ts-lethal growth defect of the readthrough mutant strain (CaDS-C, Caess1δ/Caess1TGC), we noticed that at 37°C there is a reduction in filamentation relative to the control strain (CaDS-FC; Caess1δ/CaESS1TAA) under inducing conditions. This reduction is readily apparent for cells grown for 4 days on solid Spider medium (Figure 5C, upper panels). Even after 7 days, colonies were unwrinkled compared to controls (data not shown). To confirm this result, we examined cells grown in liquid Spider medium for 2 hrs. Again, we find that the readthrough mutant (Caess1δ/Caess1TGC) is reduced in germ-tube formation relative to reconstituted control strain (Caess1v/CaESS1TAA) (Figure 5 C.). This is an intriguing result because we do not observe a growth-rate defect in the readthrough mutant when grown in standard (non-inducing) medium, even at 37°C. It seems likely therefore that the levels of Ess1 protein in this mutant strain are sufficient for standard growth but not for specialized functions such as the induction of filamentation pathways. This is consistent with findings in S. cerevisiae showing that Ess1 is present in vast excess in cells growing in standard media (YPD, CSM), but that under various stress conditions, a significant reduction in Ess1 levels rendered cells unable to grow [47].

Structure-function analysis of the C. albicans Ess1

The C. albicans Ess1 protein contains a highly-structured linker region that locks the WW and PPIase domains together [65]. Unlike the short, flexible linker in the human Pin1 protein, this linker also contains a prominent, solvent-exposed α-helix [50]. To help understand these key structural differences, we generated two mutations in the CaEss1 protein. First, we replaced residues 37–67 in C. albicans Ess1 protein with the flexible linker found in human Pin1 (residues 40–54), effectively generating a linker-swapped mutant. Second, we substituted three residues along the surface of the α-helix that would drastically alter the charge pattern it displays (E44K, A51D, and K54E; Figure 6A). As an initial test of protein function we tested their ability to complement a S. cerevisiae ts mutant (ess1H164R) at non-permissive temperature (Figure 6B) and to complement an ess1Δmutant using a plasmid-shuffle strategy (Table 1). The C. albicans linker-swapped mutant was unable to complement in either assay, whereas the helix mutant fully complemented in both assays.

Figure 6. Structure-function analysis of CaEss1.

(A) X-ray crystallographic structure of the human homolog, Pin1 [40] and CaEss1 [50]. Substitutions made in the structured linker α-helix of CaEss1 to construct the helix mutant (hm) strains are indicated. (B) Complementation of CaEss1 linker mutants in S. cerevisiae. Plasmids encoding the indicated mutant proteins (independent clone isolates of linker-swapped plasmid, pDS426(sw) and helix substitution plasmid, pDS426(pm)) were separately transformed into a ts-mutant strain of S. cerevisiae (Scess1H164R) [42]. Plasmids were constructed using a pRS426 backbone [88]. Serial dilution assays (1∶5) shows the growth of these independent transformants (a, b). Growth at 37°C, the restrictive temperature for the S. cerevisiae ess1 mutant, indicates complementing activity. The CaEss1 helix substitution mutant complements but the linker swap mutant does not. pGDCaEss1 [52] (pRS426 backbone) was used as a positive control and pRS426 [88] was used as an empty vector control. (C) Western analysis of whole cell extracts of S. cerevisiae expressing the indicated CaEss1 proteins. A total of 15 µg of protein was used per lane and the blot was probed using anti-CaEss1 polyclonal antibody at a 1∶500 dilution. For the mutant strains, two independent clone isolates of pDS413(sw) (lanes 3 and 4) and pDS413(pm) (lanes 5 and 6) were transformed into S. cerevisae and analyzed. Plasmids were constructed using a pRS413 backbone [88]. The S. cerevisiae strain used (CBW22; [44] does not express endogenous Ess1 protein but is viable due to a suppressor mutation (ess1Δsrb10Δ). The linker-swapped protein appears to be absent, or present at a very low level compared to the vector control, while the helix mutant protein is easily detected. pGD-CaESS1 (pRS413 backbone) encoding the wild-type protein (Devasahayam and Hanes, unpublished) was used as a positive control, and pRS413 was used as the empty vector control.

To confirm that the mutant C. albicans Ess1 proteins were expressed and stable in S. cerevisiae, we performed Western blot analysis (using antibodies to C. albicans Ess1). The linker-swapped mutant was present at very low levels (Figure 6C), likely explaining its failure to complement in the functional assays. Due to its presumed instability, the linker-swapped mutant was not used for further studies. In contrast, the helix mutant was expressed at much higher levels, indicating the protein is stable consistent with its functionality (Figure 6C). That the helix mutant complements S. cerevisiae ess1 mutants suggests it performs the essential catalytic function required for growth in this organism. This result is not entirely surprising given that the human and other vertebrate orthologs lacking this α-helix also rescue growth in S. cerevisiae [66]; Wilcox & Hanes, unpublished]. We interpret this result to mean either the α-helix is required solely for specialized functions, e.g. specific protein-protein interactions in C. albicans that might be lacking (or not revealed) in S. cerevisiae, or that it is not important for growth under standard conditions.

To further test the importance of this α-helix, we introduced the helix mutant (hm) alleles into C. albicans cells using the SN87 parent strain. Two strains CaDSE-5 (hm/hm) and CaDSE-5.10 (Caess1δ/hm) were constructed along with their respective isogenic controls. Both strains, CaDSE-5 and CaDSE-5.10 (Figure 1N, O) were tested for filamentation, germ tube formation and anti-fungal drug susceptibility profiles. No significant differences were seen in any of these assays (Figure 2N, O, and data not shown), suggesting the linker helix of CaEss1 is not important for morphogenetic switching or drug sensitivity in cultured cell assays. Thus, the prediction that the linker helix is important for fungal-specific functions has not yet been demonstrated. It is also possible that the triple mutation is not severe enough to inactivate its putative function(s), or that studies in animals models are needed to detect functional consequences of these mutations.

A conserved role for CaEss1 in RNA polymerase II transcription

Orthologs of CaEss1 in budding yeast (Ess1) and humans (Pin1) regulate RNA pol II transcription [42], [45], [46]. In yeast, Ess1 coordinates the recruitment of co-factors required for proper RNA synthesis and processing [43], [49]. In this role, ScEss1 is required for efficient termination of at least two classes of RNA pol II products, small non-coding RNAs, e.g. small nucleolar RNAs (snoRNAs), and protein-coding mRNAs. ScEss1 is also important for repressing the transcription of cryptic unstable transcripts (CUTs) [49]. In S. cerevisiae ess1 mutants, CUTs are detected throughout the genome, many of which are the same as those detected in RNA-decay mutants [43]. To determine whether CaEss1 plays an analogous role in C. albicans, we carried out a whole-genome transcript analysis in wild-type and CaEss1 mutant strains using high-throughput RNA sequencing (RNA-seq). The strains and conditions used are summarized in Table 2. A large number of mapped sequence reads were obtained (Table 2), indicating that our RNA libraries were of high quality.

For each mutant and control pair analyzed, there was a significant change in the level of expression for between 2–4% of all transcribed genes (Table 3). This is only slightly less than that observed in budding yeast (wild-type vs. ts-mutant) using standard microarrays where 3–10% of genes were misregulated depending on the temperature [43]. The results indicate that in C. albicans, as in S. cerevisiae the expression of some but not all genes is adversely affected by mutations in Ess1. Raw sequence reads will be deposited in the European Nucleotide Archive ( Analyzed datasets will also be submitted to the NIH Gene Expression Omnibus (GEO) ( In this report, we will focus on the comparison between the transcriptomes of the conditional (ts) readthrough mutant (ess1Δ/ess1TGC) and its isogenic control (ess1Δ/ESS1TAA), relative to the wild-type strain (ESS1/ESS1).

To determine whether CaEss1 functions in snoRNA termination, we obtained RNA-seq data from ts-mutant and control strains and examined the abundance of sequence reads at non polycistronc, independently-transcribed snoRNA genes of the box H/ACA-class [67][70]. In total, we examined expression of 26 H/ACA-class genes using Integrated Genome Viewer (IGV), and found that about one in five show evidence of readthrough as revealed by RNA-seq data. These genes included SNR3a, SNR8a, SNR32a, SNR43a, and SNR189) [67]. Primary transcripts of the other major class of snoRNAs (box C/D) are often encoded within other genes in C. albicans and were not examined here [68][70].

Each of the five H/ACA-class genes listed above contained an abundance of sequence reads (>30, but more often in the hundreds) immediately downstream of the putative termination site in the mutant background, suggesting possible transcription readthrough. The IGV profiles for three of these genes are shown (Figure 7, AC). RNA sequence reads in the forward direction (red) and reverse direction (blue) are indicated as well as the overall number of reads (grey). For the control (ess1Δ/ESS1TAA) (Figure 7AC) and wild-type (ESS1/ESS1) (data not shown) strains, the large number of reads is likely to reveal the actual position of the snoRNA transcription units (dotted grey bars) more accurately than sequence annotations (solid bars) [68][70]. The IGV also reveals examples of increased transcript abundance in the mutant cells for some snoRNAs, e.g. SNR332a, SNR43a (Figure 7B,C), and a protein-coding gene (ORF19.1968) and a non-coding transcript (CUT) (Figure 7B).

Figure 7. RNA-sequencing indicates transcription readthrough at SnoRNA loci.

Results are visualized using Integrated Genome Viewer. The total number of RNA-sequencing reads (y-axis) and their position along the chromosome (x-axis) is indicated in grey. Forward oriented reads are indicated in red; reverse oriented reads are indicated in blue. Not all reads (blue, red) are visible. Results for the CaEss1 ts-mutant strain are in the upper panels, and results for the isogenic control are in the lower panels (AC). Solid bars indicate the previously annotated gene positions, while the dotted (grey) boxes indicate the positions implied based on actual transcript data from RNA-sequencing of the the wild-type strain (ESS1/ESS1) (not shown) and the control strain (lower panels). A putative CUT is indicated in panel (B). Approximate positions of primers used for strand-specific cDNA synthesis are shown (black), as are the positions of the primer sets (green) used for qRT-PCR in (D). The positions of likely readthrough transcription are identified by red dashed arrows. (D) Results of qRT-PCR to detect readthrough transcripts for different snoRNA genes (x-axis), expressed as a fold-change (y-axis) of ts-mutant (CaDS-C) over isogenic control (CaDS-FC), normalized to ACT1. For RNA-sequencing, cell growth conditions are listed in Table 2 (shift to 42°C, then serum induction at 42°C), for qRT-PCR, samples were serum-induced at 37°C.

To confirm the presence of snoRNA readthrough transcripts, we carried out qRT-PCR (Figure 7D). cDNA synthesis was done using strand-specific primers, so only transcripts in the forward (“sense) direction relative to the snoRNA transcription units would serve as templates. Primer sets for amplification consisted of one primer within the snoRNA gene, and the other downstream of the putative termination site (Figure 7AC), therefore, only extended (readthrough) transcripts would be detected. The results show a 2–15 fold increase in the amount of readthrough transcription in the CaEss1 ts-mutant cells relative to isogenic control cells (Figure 7D). In summary, both the RNA-seq and qRT-PCR data indicate that CaEss1 is important for efficient termination of at least some snoRNA genes in C. albicans.

To determine if CaEss1 represses cryptic transcription we used IGV to compare the transcription profiles of the ts-mutant and its isogenic control, focusing on the intergenic regions along all of the chromosomes. In the mutant cells, intergenic transcription was rampant, with large amounts detected on all eight chromosomes. Examples are shown from chromomes 2, 4 and 8 (Figure 8AC). Intergenic transcripts that lie between divergent genes are probably CUTs, since they cannot be due to readthrough transcription from neighboring genes (e.g. Figure 8A). Using both short and long qRT-PCR to characterize other potential CUTs revealed that only the short products (marked with an asterisk in Figure 8B,C `) were amplified (Figure 8D, and data not shown), indicating the RNA-seq reads are not due to readthrough transcription. These data suggest that in CaEss1 mutants, cryptic promoters are activated and/or cryptic RNAs are stabilized. CUTs from each of the eight chromosomes were easily detected by qRT-PCR (e.g. Figure 8D), indicating that CUT expression is widespread in CaEss1 mutant cells.

Figure 8. RNA-sequencing reveals widespread cryptic transcription in CaEss1 ts-mutant cells.

Results are shown using Integrated Genome Viewer as described in the legend to Figure 7. The positions of cryptic unstable transcripts (CUTs) are identified on representative chromosomes (AC). (D) Results of qRT-PCR to detect CUTs (examples for each chromosome) expressed as a fold-change over wild-type (CaDS-B5) (ESS1/ESS1), normalized to ACT1. The approximate positions of the qRT-PCR products in chromosome 2, 4 and 6 samples in (D) are indicated by the short green arrows marked by an asterisk (*). The longer products indicated by long green arrows (B, C) did not amplify, indicating that the CUT signal is not likely due to a readthrough product from nearby open reading frames (ORF). cDNA synthesis was primed using a mixture of random hexamers and oligo(dT). Cell growth conditions were as listed in Table 2 (shift to 42°C, then serum induction at 42°C). (E) Prominent CUT identified within open reading frame of FGR46, which has been implicated in filamentation. (F) CUT is in reverse orientation relative to the FGR46 ORF as shown by strand-specific cDNA synthesis followed by qRT-PCR. Results are expressed as fold-change of ts-mutant (CaDS-C) over isogenic control (CaDS-FC), normalized to ACT1. Cells were serum-induced at 37°C. cDNA synthesis was primed using strand-specific primers that would reverse transcribe forward (UP044) or reverse (UP043) transcripts. For details, see Materials and Methods.

Many putative CUTs were also found within open reading frames, often in the antisense direction. One example is a potential antisense CUT within the open reading frame of FGR46 (Figure 8E), a gene implicated in filamentation by a transposon insertion screen[71]. The reverse orientation of this CUT relative to the FGR46 ORF was confirmed by strand-specific qRT-PCR (Figure 8F). Thus, it is possible that under conditions where Ess1 levels are strongly reduced, CUT induction might enhance or suppress expression of genes important for virulence. Additional analyses will be needed to fully explore the information in the RNA-sequencing datasets and determine the effect(s) of CaEss1 mutation on expression of virulence-related genes. Thus far, our findings are consistent with a conserved role for CaEss1 in transcription by RNA polymerase II.


In this study, we showed that the ESS1 gene is essential for growth in C. albicans. This result independently confirms earlier findings [52], but used more rigorous methods that included better controls and considered genomic sequence information that was not previously available. In contrast, the prior finding that reducing gene dosage by half (ess1Δ/ESS1) prevents morphogenetic switching, which is required for virulence in vivo [60], was not confirmed in the present study. The most likely explanation is that a background mutation(s) in the isolate of CAI4 used in the prior work caused the defect in filamentation. Thus, like other studies [72], [73], our results make clear the importance of reconstituted isogenic controls, appropriate marker gene systems, and the dangers of isolate-to-isolate variability between supposedly identical strains. The results of transcriptome analysis of CaEss1 mutants support the idea that regulation of RNA polymerase II by Ess1 is conserved between the evolutionarily distant species S. cerevisiae and C. albicans. Further studies will be needed to determine whether transcriptional defects are responsible for growth and morphogenetic switching phenotypes in CaEss1 mutants, and whether the mechanism by which Ess1 functions in transcription in C. albicans is similar to that in S. cerevisiae [43], [49]. With respect to C. albicans Ess1 as an antifungal drug target, data from this study suggest that elimination or strong inhibition of Ess1 enzyme activity will prevent growth. Given that Ess1 is essential not only in C. albicans, but also in A. nidulans [54], and is required for virulence in C. neoformans [53], it is possible that inhibitors of Ess1 could potentially be developed into broad-spectrum antifungal agents.

Materials and Methods

C. albicans strains, media, and transformation

All parent strains were grown in YPD media while the engineered derivative strains were grown in their respective selective media [74], unless otherwise specified. To select for URA3 loop-out events, cells were plated on YPD supplemented with 1 mg/ml 5-FOA [75]. In addition to the descriptions below, please refer to the tables for details on primers (Table 4), plasmids (Table 5) and strains (Table 6) used in this study. Transformations were performed according to the lithium acetate protocol [76].

C. abicans strain construction

Insertion of the URA3 gene at different genomic positions.

For re-evaluating the C. albicans heterozygous Caess1 mutant (CaGD1)[52] along with its control strain, R6 (CaESS1/CaESS1), the URA3 gene was removed from the CaESS1 locus using 5-FOA mediated loop-out events to create uracil auxotrophs CaGD2 (Caess1δ/CaESS1) and R6* (CaESS1/CaESS1) respectively. One copy of URA3 was integrated at the native URA3 locus by digesting pLUBP plasmid [77] with BglII and PstI restriction enzymes and transforming into the above strains (CaGD2 and R6*). Similarly, one copy of URA3 was integrated at the RPS1 locus in these strains by digesting CIp10 plasmid [78] with StuI restriction enzyme and transforming. URA3 was also integrated into the parent strain CAI4* using the same pLUBP and CIp10 plasmids. (*) indicates a strain that may have acquired one or more mutations affecting filamentation as per this study.

Isogenic wild-type control strain, CaDS-B5 (ESS1/ESS1).

Plasmid pDS426(pm) is a derivative of pRS426 that carries the CaESS1 promoter region (−255 to 0), CaESS1 ORF (with helix point mutations) and CaESS1 termination region (+535 to +735) between EcoR1 and BamH1 sites. The LEU2 marker (from plasmid pSN40) or the HIS1 marker (from plasmid pSN52) [62] was cloned into plasmid pDS426(pm) between the BamH1 and Not1 sites. These plasmids were named pDS(b)Leu and pDS(b)His, respectively. Because in the C. albicans genome, APE2 is just 210 bp downstream of ESS1, a 163 bp sequence (+572 to +735) was repeated on either side of the selectable markers to allow enough sequence for CaESS1 termination and for the 5′ APE2 promoter region. This was done by amplifying the downstream sequence of CaESS1 (+572 to +1265) with NotI ends (primers OW1075 and OW1081), digesting and inserting into pDS(b)Leu and pDS(b)His, thus positioning the NotI-NotI downstream sequence after the markers. The resulting plasmids are pDS(c)Leu and pDS(c)His.

The pDS(c)Leu and pDS(c)His plasmids were digested with SacI and serially transformed into SN87 [62]. In the first step, one CaESS1 allele was marked with the LEU2 gene to generate strain CaDS-B0.5 (ESS1: LEU2/ESS1). In the next step, the second allele of CaESS1 was marked with HIS1 gene to generate a Leu+ His+ control strain CaDS-B5 (ESS1: LEU2/ESS1: HIS1). Allele-specific PCR (primer OW221 with OW769 or OW770) and DNA sequencing was used to identify clones in which the integrating DNA recombined at positions that generated the wild-type CaESS1 ORF, (i.e. that excluded the helix mutations). The gene modifications were verified using junction-PCR and Southern blot hybridization (data not shown).

Heterozygous mutant strain, CaDS-B5.5 (ess1δ/ESS1).

The NdeI site in the backbone of the plasmid, pDS(c)His was removed by digestion with NdeI, Klenow treatment and re-ligation, to form pDS(d)His. The start codon (ATG) of CaESS1 ORF in pDS(d)His was then converted to an NdeI site with the Change-IT Multiple Mutation Site Directed Mutagenesis Kit (USB) using the mutagenesis primer OW1099. The resulting pDS(e)His plasmid was digested with Nde1 and Spe1 (site 591 bp upstream of the HIS1 marker), Klenow treated and re-ligated to generate a plasmid that lacked the ESS1 ORF sequence. The resulting plasmid, pDS(x)His was digested with SacI and transformed into strain CaDS-B0.5 (ESS1: LEU2/ESS1) to generate a heterozygous mutant, CaDS-B5.5 (ESS1: LEU2/ess1δ: HIS1). The gene modifications were verified using junction-PCR and Southern blot hybridization (data not shown). In plasmid, pDS(x)His the HIS1 marker (from the pSN52 plasmid) was left with a 591 bp promoter region. An additional 393 bp sequence that was upstream of the promoter region of HIS1 is removed in this construct. This had no affect on HIS1 expression by qRT-PCR (primers OW1415 and D25) in comparison to strain CaDS-B5 (ESS1/ESS1) (data not shown).

Temperature-sensitive strain, CaDS-C (ess1δ/ess1(TGC) and isogenic control.

The plasmid pDS(x)His was digested with SacI and transformed into SN87 to generate CaDS-87-2.3 (ess1δ/ESS1), which is auxotrophic for LEU2. The Nde1 site of the vector portion in plasmid pDS(c)Leu was removed (as done for pDS(c)His), to form pDS(d)Leu. Next, the start codon (ATG) and the stop codon (TAA) of CaESS1 in plasmid pDS(d)Leu were changed to an NdeI site and a SphI site (primers OW1099 and OW1100, respectively) in two steps, to first form plasmid pDS(e)Leu and then plasmid pDS(f)Leu, respectively. The wild-type CaESS1 sequence was amplified (primers OW1354 and OW1355) with flanking Nde1 and Sph1 sites, digested and inserted into the same sites of pDS(f)Leu to give pDS(g)Leu. The Sph1 site changes the stop codon from TAA to TGC. The pDS(g)Leu plasmid was digested with SacI and transformed into CaDS87-2.3 (ess1δ/ESS1) to make a prototrophic strain, CaDS-C (ess1δ/ess1TGC). This strain was verified using junction-PCR and Southern blot hybridization (data not shown). The control strain was generated by a novel strategy. About a 900 bp PCR fragment amplified (using primers OW216 and OW1231) from a wild-type background containing the normal termination codon (TAA) was introduced into CaDS-C (ess1δ/Caess1TGC). Transformants capable of growth at 42°C were selected, analyzed by PCR amplification and DNA sequencing of the CaESS1 alleles and the mutation of TGC to TAA was confirmed. The resulting prototrophic strain CaDS-FC (ess1δ/ESS1TAA) produced Ess1 protein of the normal size (19 kDa) and was non-ts.

Linker-helix mutations in CaEss1.

PCR overlap extension (with primer sets OW748-OW1224, OW749-OW1233, and OW1224-OW1233) was used to generate a fragment that encodes CaEss1 protein bearing the linker region of human Pin1 (codon optimized for C. albicans [79]. This linker-swap fragment was digested with EcoRI and BamHI sites and cloned into the same sites of pRS426 vector to form pDS426(sw). A similar strategy was used to generate a CaEss1 mutant bearing three amino acid substitutions (E44K, A51D, and K54E) in the linker helix (using primers OW750 and OW751). The helix mutant fragment was cloned into pRS426 vector to form pDS426(pm).

Helix mutants, CaDS-E (ESS1/hm), CaDS-E5 (hm/hm), and CaDS-E5.10 (ess1δ/hm).

To construct CaDS-E (ESS1/hm) strains, the plasmid, pDS(c)Leu was digested with SacI and transformed into SN87. Allele-specific PCR and DNA sequencing was used to verify the clones that recombined to include the helix mutations (Caess1hm). This strain is auxotrophic for HIS1. To construct prototrophic CaDS-E5 (hm/hm) strains, pDS(c)His was digested with SacI and transformed into CaDS-E (ESS1/hm). To construct CaDS-E5.10 (ess1δ/hm) strains, pDS(x)His was digested with SacI and transformed into CaDS-E(ESS1/hm). Mutants were verified using junction-PCR and Southern blot hybridization (data not shown).

APE2 promoter mutant, CaDS-51B-2 (5′ape2/5′ape2).

A Not1-Not1 PCR fragment (primers D31 and OW1081) was cloned into plasmids pDS(b)Leu and pDS(b)His, to form plasmids pDS(h)Leu and pDS(h)His, respectively. This fragment (+729 to +1265) contained only 15 bp of 5′ genomic sequence upstream of APE2 (and an upstream Not1 site). The plasmid was digested with SacI and transformed into SN87, in two steps to generate strain CaDS-51B-2 (5ape2/5′ape2).

Growth assays

Overnight cultures of either S. cerevisiae or C. albicans were diluted to an OD600 of 0.1 and grown with shaking at 30°C until midlog phase. Cultures were brought to an OD600 of 0.5 either by dilution or concentration by centrifugation. For S. cerevisiae, a 1∶5 dilution series of the strains were spotted onto plates containing selective media and incubated for two nights. For C. albicans strains, a 1∶3 dilution series was used, with cells spotted on YPD plates and incubated overnight.

Filamentation assays

Overnight cultures of C. albicans were diluted to an OD600 of 0.25 (∼3×106 cells/ml). From dilution, 2 µl of culture was spotted onto Spider media [80] and serum-containing media (4% FBS and 2% agar). The plates were incubated at 37°C for 4 days prior to being photographed. To test germ tube formation, a suspension of 105–106 cells/ml was made in liquid Spider medium [80], and incubated at 37°C for 2 hrs prior to microscopic observation.

Western blot analysis

Overnight cultures were diluted to an OD600 of 0.1, grown to an OD600 of 0.5 and harvested by centrifugation. Protein extracts were prepared using yeast protein lysis buffer (50 mM HEPES, 140 mM NaCl, 1 mM EDTA, 1% Triton X and, 0.1% sodium deoxycholate) with proteinase inhibitor cocktail (ethanolic protease inhibitors with 0.25 M PMSF and 0.7 mg/ml pepstatin/aqueous protease inhibitors with 0.1 mg/ml leupeptin, 1 mg/ml soybean trypsin inhibitor, 0.1 mg/ml aprotinin in 10 mM Tris, pH7.5, 20 mM benzamidine, 10 mM sodium vanadate and, 500 mM sodium fluoride). A 15% polyacrylamide gel was run at 100 V for about 2.5 hrs and transferred to a PVDF Transfer Membrane (Millipore) overnight at 32 V at 4°C. A 1∶500 dilution of rabbit α-CaEss1 polyclonal antibodies (Applied Biosystems) was used as primary antibody and a 1∶25,000 dilution of horseradish peroxidase-linked donkey anti-rabbit IgG antibody (GE Healthcare) was used as the secondary antibody. Purified CaEss1 made in E. coli [50] was used to generate CaEss1 polyclonal antibodies at Applied Biosystems. Detection was done using the ECL-Plus Western Blotting Detection System (GE Healthcare) and exposure to X-ray film (Kodak).

High-throughput RNA sequencing

RNA and protein preparation.

For the ts-readthrough mutant and control strain CaDS-C (Caess1δ/Caess1TGC) and CaDS-FC (Caess1δ/CaESS1TAA) the following protocol was used. Cultures (10 ml) were grown overnight in selective media at 30°C, diluted 10-fold in YPD, and grown overnight again to an OD600 of 9–11 [81]. Cultures were diluted 30 times in pre-warmed (42°C), pre-shaken fresh YPD (1 ml in 29 ml YPD)[81], and cells were grown at 42°C with shaking. After 1 hr, 2 ml of culture was collected for protein preparation as above and the remaining culture diluted 2-fold in pre-warmed (42°C), pre-shaken YPD+20% FBS (20 ml culture in 20 ml YPD+20% FBS) for serum induction. After 2 hr incubation at 42°C, cultures were collected by centrifugation for protein preparation (2 ml) and RNA extraction (10 ml). At extraction the cells were at an OD600 of 0.4. RNA was extracted by the hot phenol method [82]. For all other strains the following protocol used. Overnight cultures in YPD were grown to an OD600 of 9–11, and diluted 20-fold in pre-warmed (37°C), pre-shaken fresh YPD+10% FBS (1 ml culture in 19 ml YPD+10% FBS) [81]. After a 2 hr incubation at 37°C, 10 ml of culture (OD600 of 0.8) was collected for RNA extraction.

RNA was quantified using a Nanodrop Spectrophotometer and aliquots of 10 µg of RNA subjected to DNase digestion (Epicentre) for 45 min at 37°C. Digested RNA was purified using an RNeasy Mini Kit (Qiagen). 400 ng of RNA was fractionated by agarose (1%) gel electrophoresis to confirm integrity (data not shown). For the ts-readthrough strain and its isogenic control, protein extracts before and after the temperature shift and serum induction were analyzed by Western blot analysis to confirm reduced amount of Ess1 protein at the restrictive temperature (42°C).

Library preparation and high-throughput sequencing.

RNA samples (1 µg in 10 µl of RNase free water) were sent for library preparation and sequencing to off-site facilities (Weill Cornell Medical College, NY and Centrillion Biosciences, CA). At Cornell, from 1 µg of purified RNA, ribosomal RNA was removed using Ribo-Zero rRNA Removal Kit (Epicenter). Illumina-compatible, barcoded, strand-specific, cDNA libraries were prepared using ScriptSeq mRNA-Seq Library Preparation Kit and RNA-Seq Barcode Primers (Epicenter). Illumina high-throughput sequencing was performed on the libraries using the HiSeq2000 sequencer running at 58 cycles (per lane) resulting in one-end read lengths of about 51 bp. At Centrillion, a similar approach was taken to prepare Illumina-compatible, barcoded, strand-specific, cDNA libraries with the exceptions of using the RiboMinus Eukaryote Kit (Invitrogen) for removal of ribosomal RNA and the ScriptSeq V2 RNA-Seq Library Preparation Kit (Epicenter). The ScriptSeq V2 Kit is an improved version of the ScriptSeq Kit and uses the same basic principal for library construction. Here the HiSeq2000 sequencer was run at 58×2 cycles (per lane) resulting paired-end read lengths of about 75 bp.

RNA sequence analysis.

The raw FastQ data files were aligned to the C. albicans genome (Assembly 21) [83] obtained from Candida Genome Database ( using Bowtie/TopHat short sequence read alignment software programs [84], [85]. Each strain resulted in a large number of reads mapped to the genome (Table 2). The output files (in SAM format) were converted to BAM files using SAM Tools [86]. The BAM files were visually analyzed on Integrative Genomics Viewer ( and/or GenomeView ( The files were quantitatively analyzed using the Galaxy server (, an open access platform for high-throughput data analysis. For example, the SAM or BAM files were used as Cuffdiff (a part of the open-source Cufflinks software package) [85] input files to identify differential expression of transcripts genome-wide among test strains and controls.

Quantitative reverse transcription real-time PCR

Overnight cultures were diluted to an OD600 of 0.5 and grown to an OD600 of 0.8–1.2 and harvested. RNA was extracted using the hot phenol method [82] and subjected to DNase digestion (Epicenter) for 45 min at 37°C. From 1 µg of RNA, cDNA was synthesized using the First Strand cDNA Synthesis Kit for Real-Time PCR(USB) using mixture of oligo dT and random hexamers. In some sets of experiments cDNA was synthesized using individual cDNA synthesis reagents purchased from USB/Affimetrix. Quantitative real-time PCR was performed in the ABI2000 RT-PCR machine using the HotStart-IT SYBR Green qPCR Master Mix (USB). In some sets of experiments real-time PCR was carried out using the Fermentus SYBR Green Master Mix (Life Sciences). All results were normalized agains the same internal control gene, ACT1. qRT-PCR calculations were done as per Yu et al., [87] where the quantitative real-time PCR cycle numbers relative to ACT1 were summed for each biological replicate and normalized to the summed average of all samples within a given experiment. The re-normalized values were then used for average and standard deviation calculations. This method corrects for trial-to-trial variability. To calculate the fold difference between samples, the ACT1-normalized (ΔCt) values for the FC (WT/-) and C (ts/−) data sets were normalized against the WT/WT (B5) data (to obtain ΔΔCt), and the following calculation was performed: fold difference = 2−ΔΔCt. The values obtained were averaged, and a standard deviation calculated.


We are grateful to Susanne Noble, William Fonzi and Alistair Brown for strains and plasmids, to Daniel Diekema for conducting the anti-fungal susceptibility assays, to Mike Palumbo for help with the bioinformatics analysis, to Navjot Singh for helpful discussions and comments on the manuscript, and to the Wadsworth Center Molecular Genetics, Microarray, Bioinformatics, and Media Core Facilities.

Author Contributions

Conceived and designed the experiments: DS DA RM JTW VC SDH. Performed the experiments: DS DA. Analyzed the data: DS DA RM JTW VC SDH. Contributed reagents/materials/analysis tools: DS DA RM JTW VC SDH. Wrote the paper: DS SDH.


  1. 1. Schmid J, Tortorano AM, Jones G, Lazzarini C, Zhang N, et al. (2011) Increased mortality in young candidemia patients associated with presence of a Candida albicans general-purpose genotype. J Clinical Microbiol 49: 3250–3256.
  2. 2. Mean M, Marchetti O, Calandra T (2008) Bench-to-bedside review: Candida infections in the intensive care unit. Critical Care 12: 204.
  3. 3. Pfaller MA, Messer SA, Moet GJ, Jones RN, Castanheira M (2011) Candida bloodstream infections: comparison of species distribution and resistance to echinocandin and azole antifungal agents in Intensive Care Unit (ICU) and non-ICU settings in the SENTRY Antimicrobial Surveillance Program. 2008–2009Intern J Antimicrobial Agents 38: 65–69.
  4. 4. Yang YL, Leaw SN, Wang AH, Chen HT, Cheng WT, et al. (2011) Characterization of yeasts colonizing in healthy individuals. Medical Mycology 49: 103–106.
  5. 5. Gow NA, van de Veerdonk FL, Brown AJ, Netea MG (2012) Candida albicans morphogenesis and host defence: discriminating invasion from colonization. Nat Rev Microbiol 10: 112–122.
  6. 6. Odds FC (1994) Candida albicans, the life and times of a pathogenic yeast. J Med Vet Mycol 32 Suppl 1: 1–8.
  7. 7. Odds FC (1994) Pathogenesis of Candida infections. J Am Acad Derm 31: S2–5.
  8. 8. Mavor AL, Thewes S, Hube B (2005) Systemic fungal infections caused by Candida species: epidemiology, infection process and virulence attributes. Current Drug Targets 6: 863–874.
  9. 9. De Rosa FG, Garazzino S, Pasero D, Di Perri G, Ranieri VM (2009) Invasive candidiasis and candidemia: new guidelines. Minerva Anestesiologica 75: 453–458.
  10. 10. Lai CC, Tan CK, Huang YT, Shao PL, Hsueh PR (2008) Current challenges in the management of invasive fungal infections. J Infect Chemo 14: 77–85.
  11. 11. Schelenz S, Abdallah S, Gray G, Stubbings H, Gow I, et al. (2011) Epidemiology of oral yeast colonization and infection in patients with hematological malignancies, head neck and solid tumors. J Oral Path Med 40: 83–89.
  12. 12. Epstein JB, Hancock PJ, Nantel S (2003) Oral candidiasis in hematopoietic cell transplantation patients: an outcome-based analysis. Oral Surg Med Path Rad Endo 96: 154–163.
  13. 13. Muller FM, Groll AH, Walsh TJ (1999) Current approaches to diagnosis and treatment of fungal infections in children infected with human immuno deficiency virus. Eur J Pediatrics 158: 187–199.
  14. 14. Fridkin SK, Kaufman D, Edwards JR, Shetty S, Horan T (2006) Changing incidence of Candida bloodstream infections among NICU patients in the United States. 1995–2004Pediatrics 117: 1680–1687.
  15. 15. Cassone A, Cauda R (2012) Candida and candidiasis in HIV-infected patients: where commensalism, opportunistic behavior and frank pathogenicity lose their borders. AIDS 26: 1457–1472.
  16. 16. Jain A, Jain S, Rawat S (2010) Emerging fungal infections among children: A review on its clinical manifestations, diagnosis, and prevention. J Pharm Bioallied Sci 2: 314–320.
  17. 17. Casalinuovo IA, Di Francesco P, Garaci E (2004) Fluconazole resistance in Candida albicans: a review of mechanisms. Eur Rev Med Pharmacol Sci 8: 69–77.
  18. 18. Cannon RD, Lamping E, Holmes AR, Niimi K, Tanabe K, et al. (2007) Candida albicans drug resistance another way to cope with stress. Microbiology 153: 3211–3217.
  19. 19. Pitman SK, Drew RH, Perfect JR (2011) Addressing current medical needs in invasive fungal infection prevention and treatment with new antifungal agents, strategies and formulations. Expert Opin Emerging Drugs. Epub Aug 17.
  20. 20. Huang M, Kao KC (2012) May 21 Population dynamics and the evolution of antifungal drug resistance in Candida albicans. FEMS Micro Let epub
  21. 21. Pfaller MA, Moet GJ, Messer SA, Jones RN, Castanheira M (2011) Candida bloodstream infections: comparison of species distributions and antifungal resistance patterns in community-onset and nosocomial isolates in the SENTRY Antimicrobial Surveillance Program. 2008–2009Antimicrobial Agents Chemo 55: 561–566.
  22. 22. Vazquez JA (2003) Combination antifungal therapy against Candida species: the new frontier-are we there yet? Medical Mycology 41: 355–368.
  23. 23. Kontoyiannis DP, Lewis RE (2004) Toward more effective antifungal therapy: the prospects of combination therapy. British J Haematol 126: 165–175.
  24. 24. Agarwal AK, Tripathi SK, Xu T, Jacob MR, Li XC, et al. (2012) Exploring the molecular basis of antifungal synergies using genome-wide approaches. Frontiers Micro 3: 115.
  25. 25. Garbati MA, Alasmari FA, Al-Tannir MA, Tleyjeh IM (2012) The role of combination antifungal therapy in the treatment of invasive aspergillosis: a systematic review. International J Infect Diseases 16: e76–81.
  26. 26. Del Pozo JL, Frances ML, Hernaez S, Serrera A, Alonso M, et al. (2011) Effect of amphotericin B alone or in combination with rifampicin or clarithromycin against Candida species biofilms. Intern J Artificial Organs 34: 766–770.
  27. 27. Mahboubi M, Ghazian Bidgoli F (2010) In vitro synergistic efficacy of combination of amphotericin B with Myrtus communis essential oil against clinical isolates of Candida albicans. Phytomedicine 17: 771–774.
  28. 28. Edlich F, Fischer G (2006) Pharmacological targeting of catalyzed protein folding: the example of peptide bond cis/trans isomerases. Handbook of Exp Pharma 359–404.
  29. 29. Bell A, Monaghan P, Page AP (2006) Peptidyl-prolyl cis-trans isomerases (immunophilins) and their roles in parasite biochemistry, host-parasite interaction and antiparasitic drug action. Intern J Parasit 36: 261–276.
  30. 30. Schiene C, Fischer G (2000) Enzymes that catalyse the restructuring of proteins. Curr Opin Struct Biol 10: 40–45.
  31. 31. Schiene-Fischer C, Aumuller T, Fischer G (2012) May 20 Peptide Bond cis/trans Isomerases: A Biocatalysis Perspective of Conformational Dynamics in Proteins. Topics in Curr Chem [Epub ahead of print] PMID 21598101.
  32. 32. Arevalo-Rodriguez M, Wu X, Hanes SD, Heitman J (2004) Prolyl isomerases in yeast. Front Biosci 9: 2420–2446.
  33. 33. Derkx PM, Madrid SM (2001) The Aspergillus niger cypA gene encodes a cyclophilin that mediates sensitivity to the immunosuppressant cyclosporin A. Mol Genet Genomics 266: 527–536.
  34. 34. Cruz MC, Goldstein AL, Blankenship J, Del Poeta M, Perfect JR, et al. (2001) Rapamycin and less immunosuppressive analogs are toxic to Candida albicans and Cryptococcus neoformans via FKBP12-dependent inhibition of TOR. Antimicrobial Agents and Chemo 45: 3162–3170.
  35. 35. Cruz MC, Cavallo LM, Gorlach JM, Cox G, Perfect JR, et al. (1999) Rapamycin antifungal action is mediated via conserved complexes with FKBP12 and TOR kinase homologs in Cryptococcus neoformans. Mol Cell Biol 19: 4101–4112.
  36. 36. Bastidas RJ, Reedy JL, Morales-Johansson H, Heitman J, Cardenas ME (2008) Signaling cascades as drug targets in model and pathogenic fungi. Curr Opin Investig Drugs 9: 856–864.
  37. 37. Baumgrass R, Zhang Y, Erdmann F, Thiel A, Weiwad M, et al. (2004) Substitution in position 3 of cyclosporin A abolishes the cyclophilin-mediated gain-of-function mechanism but not immunosuppression. J Biol Chem 279: 2470–2479.
  38. 38. Singh N, Heitman J (2004) Antifungal attributes of immunosuppressive agents: new paradigms in management and elucidating the pathophysiologic basis of opportunistic mycoses in organ transplant recipients. Transplantation 77: 795–800.
  39. 39. Hani J, Stumpf G, Domdey H (1995) PTF1 encodes an essential protein in Saccharomyces cerevisiae, which shows strong homology with a new putative family of PPIases. FEBS Lett 365: 198–202.
  40. 40. Ranganathan R, Lu KP, Hunter T, Noel JP (1997) Structural and functional analysis of the mitotic rotamase Pin1 suggests substrate recognition is phosphorylation dependent. Cell 89: 875–886.
  41. 41. Hanes SD, Shank PR, Bostian KA (1989) Sequence and mutational analysis of ESS1, a gene essential for growth in Saccharomyces cerevisiae. Yeast 5: 55–72.
  42. 42. Wu X, Wilcox CB, Devasahayam G, Hackett RL, Arevalo-Rodriguez M, et al. (2000) The Ess1 prolyl isomerase is linked to chromatin remodeling complexes and the general transcription machinery. EMBO J 19: 3727–3738.
  43. 43. Singh N, Ma Z, Gemmill T, Wu X, Rossettini A, Rabeler C, Beane O, DeFiglio H, Palumbo M, Morse R, Hanes, S D (2009) The Ess1 prolyl isomerase is required for transcription termination of small non-coding regulatory RNAs via the Nrd1 pathway. Mol Cell 36: 255–266.
  44. 44. Wilcox CB, Rossettini A, Hanes SD (2004) Genetic interactions with C-terminal domain (CTD) kinases and the CTD of RNA Pol II suggest a role for Ess1 in transcription initiation and elongation in Saccharomyces cerevisiae. Genetics 167: 93–105.
  45. 45. Xu YX, Hirose Y, Zhou XZ, Lu KP, Manley JL (2003) Pin1 modulates the structure and function of human RNA polymerase II. Genes Dev 17: 2765–2776.
  46. 46. Krishnamurthy S, Ghazy MA, Moore C, Hampsey M (2009) Functional interaction of the Ess1 prolyl isomerase with components of the RNA polymerase II initiation and termination machineries. Mol Cell Biol 29: 2925–2934.
  47. 47. Gemmill TR, Wu X, Hanes SD (2005) Vanishingly low levels of Ess1 prolyl-isomerase activity are sufficient for growth in Saccharomyces cerevisiae. J Biol Chem 280: 15510–15517.
  48. 48. Morris DP, Phatnani HP, Greenleaf AL (1999) Phospho-carboxyl-terminal domain binding and the role of a prolyl isomerase in pre-mRNA 3′-End formation. J Biol Chem 274: 31583–31587.
  49. 49. Ma Z, Atencio D, Barnes C, DeFiglio H, Hanes SD (2012) Multiple Roles for the Ess1 Prolyl Isomerase in the RNA Polymerase II Transcription Cycle. Mol Cell Biol 32: 3594–3607.
  50. 50. Li Z, Li H, Devasahayam G, Gemmill T, Chaturvedi V, et al. (2005) The structure of the Candida albicans Ess1 prolyl isomerase reveals a well-ordered linker that restricts domain mobility. Biochemistry 44: 6180–6189.
  51. 51. Verdecia MA, Bowman ME, Lu KP, Hunter T, Noel JP (2000) Structural basis for phosphoserine-proline recognition by group IV WW domains. Nat Struct Biol 7: 639–643.
  52. 52. Devasahayam G, Chaturvedi V, Hanes SD (2002) The Ess1 prolyl isomerase is required for growth and morphogenetic switching in Candida albicans. Genetics 160: 37–48.
  53. 53. Ren P, Rossettini A, Chaturvedi V, Hanes SD (2005) The Ess1 prolyl isomerase is dispensable for growth but required for virulence in Cryptococcus neoformans. Microbiology 151: 1593–1605.
  54. 54. Joseph JD, Daigle SN, Means AR (2004) PINA is essential for growth and positively influences NIMA function in Aspergillus nidulans. J Biol Chem 279: 32373–32384.
  55. 55. Fonzi WA, Irwin MY (1993) Isogenic strain construction and gene mapping in Candida albicans. Genetics 134: 717–728.
  56. 56. Lay J, Henry LK, Clifford J, Koltin Y, Bulawa CE, et al. (1998) Altered expression of selectable marker URA3 in gene-disrupted Candida albicans strains complicates interpretation of virulence studies. Infect Immun 66: 5301–5306.
  57. 57. Brand A, MacCallum DM, Brown AJ, Gow NA, Odds FC (2004) Ectopic expression of URA3 can influence the virulence phenotypes and proteome of Candida albicans but can be overcome by targeted reintegration of URA3 at the RPS10 locus. Euk Cell 3: 900–909.
  58. 58. Magee PT, Gale C, Berman J, Davis D (2003) Molecular genetic and genomic approaches to the study of medically important fungi. Infect Infect Immun 71: 2299–2309.
  59. 59. Jones T, Federspiel NA, Chibana H, Dungan J, Kalman S, et al. (2004) The diploid genome sequence of Candida albicans. Proc Natl Acad Sci USA 101: 7329–7334.
  60. 60. Sanchez AA, Johnston DA, Myers C, Edwards JE Jr, Mitchell AP, et al. (2004) Relationship between Candida albicans virulence during experimental hematogenously disseminated infection and endothelial cell damage in vitro. Infect Immun 72: 598–601.
  61. 61. Klinke T, Rump A, Ponisch R, Schellenberger W, Muller EC, et al. (2008) Identification and characterization of CaApe2-a neutral arginine/alanine/leucine-specific metallo-aminopeptidase from Candida albicans. FEMS Yeast Res 8: 858–869.
  62. 62. Noble SM, Johnson AD (2005) Strains and strategies for large-scale gene deletion studies of the diploid human fungal pathogen Candida albicans. Euk Cell 4: 298–309.
  63. 63. Moreno I, Martinez-Esparza M, Laforet LC, Sentandreu R, Ernst JF, et al. (2010) Dosage-dependent roles of the Cwt1 transcription factor for cell wall architecture, morphogenesis, drug sensitivity and virulence in Candida albicans. Yeast 27: 77–87.
  64. 64. Porta A, Ramon AM, Fonzi WA (1999) PRR1, a homolog of Aspergillus nidulans palF, controls pH-dependent gene expression and filamentation in Candida albicans. J Bact 181: 7516–7523.
  65. 65. McNaughton L, Li Z, Van Roey P, Hanes SD, LeMaster DM (2010) Restricted domain mobility in the Candida albicans Ess1 prolyl isomerase. Biochimica et biophysica acta 1804: 1537–1541.
  66. 66. Lu KP, Hanes SD, Hunter T (1996) A human peptidyl-prolyl isomerase essential for regulation of mitosis. Nature 380: 544–547.
  67. 67. Mitrovich QM, Tuch BB, De La Vega FM, Guthrie C, Johnson AD (2010) Evolution of yeast noncoding RNAs reveals an alternative mechanism for widespread intron loss. Science 330: 838–841.
  68. 68. Sellam A, Hogues H, Askew C, Tebbji F, van Het Hoog M, et al. (2010) Experimental annotation of the human pathogen Candida albicans coding and noncoding transcribed regions using high-resolution tiling arrays. Genome Biol 11: R71.
  69. 69. Schattner P, Decatur WA, Davis CA, Ares M Jr, Fournier MJ, et al. (2004) Genome-wide searching for pseudouridylation guide snoRNAs: analysis of the Saccharomyces cerevisiae genome. Nuc Acids Res 32: 4281–4296.
  70. 70. Lowe TM, Eddy SR (1999) A computational screen for methylation guide snoRNAs in yeast. Science 283: 1168–1171.
  71. 71. Uhl MA, Biery M, Craig N, Johnson AD (2003) Haploinsufficiency-based large-scale forward genetic analysis of filamentous growth in the diploid human fungal pathogen C. albicans. EMBO J 22: 2668–2678.
  72. 72. Cheng S, Nguyen MH, Zhang Z, Jia H, Handfield M, et al. (2003) Evaluation of the roles of four Candida albicans genes in virulence by using gene disruption strains that express URA3 from the native locus. Infec Imm 71: 6101–6103.
  73. 73. Sundstrom P, Cutler JE, Staab JF (2002) Reevaluation of the role of HWP1 in systemic candidiasis by use of Candida albicans strains with selectable marker URA3 targeted to the ENO1 locus. Infec Imm 70: 3281–3283.
  74. 74. Sherman F (1991) Getting started with Yeast, Ed Guthrie, C. and Fink, G. R. Meth in Enzymology 194: 1–21.
  75. 75. Wellington M, Rustchenko E (2005) 5-Fluoro-orotic acid induces chromosome alterations in Candida albicans. Yeast 22: 57–70.
  76. 76. Ito H, Fukuda Y, Murata K, Kimura A (1983) Transformation of intact yeast cells treated with alkali cations. J. Bact 153: 163–168.
  77. 77. Ramon AM, Fonzi WA (2003) Diverged binding specificity of Rim101p, the Candida albicans ortholog of PacC. Eukaryotic Cell 2: 718–728.
  78. 78. Murad AM, Lee PR, Broadbent ID, Barelle CJ, Brown AJ (2000) CIp10, an efficient and convenient integrating vector for Candida albicans. Yeast 16: 325–327.
  79. 79. Lloyd AT, Sharp PM (1992) Evolution of codon usage patterns: the extent and nature of divergence between Candida albicans and Saccharomyces cerevisiae. Nuc Acids Res 20: 5289–5295.
  80. 80. Liu H, Kohler J, Fink GR (1994) Suppression of hyphal formation in Candida albicans by mutation of a STE12 homolog. Science 266: 1723–1726.
  81. 81. Kadosh D, Johnson AD (2005) Induction of the Candida albicans filamentous growth program by relief of transcriptional repression: a genome-wide analysis. Mol Biol Cell 16: 2903–2912.
  82. 82. Schmitt ME, Brown TA, Trumpower BL (1990) A rapid and simple method for preparation of RNA from Saccharomyces cerevisiae. Nuc Acids Res 18: 3091–3092.
  83. 83. van het Hoog M, Rast TJ, Martchenko M, Grindle S, Dignard D, et al. (2007) Assembly of the Candida albicans genome into sixteen supercontigs aligned on the eight chromosomes. Genome Biol 8: R52.
  84. 84. Langmead B, Trapnell C, Pop M, Salzberg SL (2009) Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol 10: R25.
  85. 85. Trapnell C, Pachter L, Salzberg SL (2009) TopHat: discovering splice junctions with RNA-Seq. Bioinformatics 250: 1105–1111.
  86. 86. Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, et al. (2009) The Sequence Alignment/Map format and SAMtools. Bioinformatics 25: 2078–2079.
  87. 87. Yu C, Palumbo MJ, Lawrence CE, Morse RH (2006) Contribution of the histone H3 and H4 amino termini to Gcn4p- and Gcn5p-mediated transcription in yeast. J Biol Chem 281: 9755–9764.
  88. 88. Sikorski RS, Hieter P (1989) A system of shuttle vectors and yeast host strains designed for efficient manipulation of DNA in Saccharomyces cerevisiae. Genetics 122: 19–27.