The SAGA and NuA4 component Tra1 regulates Candida albicans drug resistance and pathogenesis

Abstract

Candida albicans is the most common cause of death from fungal infections. The emergence of resistant strains reducing the efficacy of first-line therapy with echinocandins, such as caspofungin calls for the identification of alternative therapeutic strategies. Tra1 is an essential component of the SAGA and NuA4 transcriptional co-activator complexes. As a PIKK family member, Tra1 is characterized by a C-terminal phosphoinositide 3-kinase domain. In Saccharomyces cerevisiae, the assembly and function of SAGA and NuA4 are compromised by a Tra1 variant (Tra1_Q3) with three arginine residues in the putative ATP-binding cleft changed to glutamine. Whole transcriptome analysis of the S. cerevisiae tra1_Q3 strain highlights Tra1’s role in global transcription, stress response, and cell wall integrity. As a result, tra1_Q3 increases susceptibility to multiple stressors, including caspofungin. Moreover, the same tra1_Q3 allele in the pathogenic yeast C. albicans causes similar phenotypes, suggesting that Tra1 broadly mediates the antifungal response across yeast species. Transcriptional profiling in C. albicans identified 68 genes that were differentially expressed when the tra1_Q3 strain was treated with caspofungin, as compared to gene expression changes induced by either tra1_Q3 or caspofungin alone. Included in this set were genes involved in cell wall maintenance, adhesion, and filamentous growth. Indeed, the tra1_Q3 allele reduces filamentation and other pathogenesis traits in C. albicans. Thus, Tra1 emerges as a promising therapeutic target for fungal infections.

fungal genetics, fungal pathogenesis, drug resistance, Candida, SAGA, stress response

Introduction

Fungal infections are a major modern public health challenge, killing over one million people annually (Brown et al. 2012; Bongomin et al. 2017). While the human immune system presents an effective barrier to infection in healthy individuals, most fungal pathogens are opportunistic and can cause deadly invasive infections in immunosuppressed patients. Candida species are amongst the most common causes of life-threatening invasive fungal infections, with Candida albicans being the most frequently isolated pathogen (Pfaller and Diekema 2007; Kullberg and Arendrup 2015). Currently, there is a limited armamentarium of effective, nontoxic antifungal therapeutics for the treatment of invasive candidiasis, with three major classes of antifungal drugs in clinical use: polyenes, azoles, and echinocandins (Perfect 2017). However, acquired resistance to these antifungal agents is increasingly common amongst C. albicans isolates, and non-albicans Candida species with high rates of acquired or intrinsic antifungal resistance (Cowen et al. 2002; Lee et al. 2021), including the emerging multidrug-resistant pathogen Candida auris, becoming increasingly prevalent (Miceli et al. 2011; Whaley et al. 2016; Chowdhary et al. 2017; Geddes-McAlister and Shapiro 2019).

The echinocandins, which include caspofungin, are the most recently discovered class of antifungal drugs and are fungicidal against most Candida pathogens (Aguilar-Zapata et al. 2015). Echinocandins cause significant cell wall stress by inhibiting the (1,3)-β-d-glucan synthase (encoded in Candida by FKS1/2), and thus decrease production of the critical cell wall component (1,3)-β-d-glucan (Kurtz and Douglas 1997; Pound et al. 2010). In many cases, echinocandins are the first line of treatment for invasive candidiasis, and the majority of patients with candidemia receive echinocandins (Cleveland et al. 2012; Perlin 2015; Pappas et al. 2016). As a result, a growing proportion of disease-causing Candida isolates display reduced susceptibility to echinocandins, and in some cases, cross-resistance with other antifungals such as azoles (Garcia-Effron et al. 2010; Alexander et al. 2013; Perlin 2015). The increasing threat of antifungal resistance, in combination with the already limited diversity of antifungal drug classes, make it imperative to discover alternative lines of treatment and expand the availability of effective therapeutic approaches.

Antifungal drugs such as the echinocandins activate fungal stress responses, which are important in the susceptibility and resistance to antifungal therapeutics (Cannon et al. 2007; Cowen and Steinbach 2008; Shor and Perlin 2015; Víglaš and Olejníková 2020). As with other stress responses, these antifungal stress response pathways are controlled through gene expression. In eukaryotic organisms, chromatin modifications are critical for the regulation of gene expression (Bannister and Kouzarides 2011; Yun et al. 2011). One role of modifications, such as the acetylation of lysine residues on the N-terminal tails of histones, is to facilitate the recruitment of the transcriptional machinery to target promoters. The SAGA (Spt-Ada-Gcn5-Acetyltransferase) and NuA4 (Nucleosome acetyltransferase of H4) complexes contain two of the principal lysine acetyltransferases, Gcn5, and Esa1, respectively (Grant et al. 1997; Allard et al. 1999; Auger et al. 2008). Both target lysines within the N-terminal tails of histones and have substrates with roles unrelated to chromatin (Choudhary et al. 2009; Henriksen et al. 2012; Mitchell et al. 2013). The SAGA complex has a broad role in transcription, including regulating multiple stress response pathways (Huisinga and Pugh 2004; Bonnet et al. 2014; Baptista et al. 2018).

Tra1/TRRAP is an essential component of SAGA and NuA4 complexes (Grant et al. 1998; Saleh et al. 1998; Elías-Villalobos et al. 2019b). It is a member of the PIKK (phosphoinositide-3-kinase-related kinase) family that also includes Tor1, ATM and Rad3-related protein (ATR/Mec1), and the DNA-dependent protein kinase catalytic subunit (DNA-PKcs) (Lempiäinen and Halazonetis 2009). As with other PIKKs, Tra1 is incorporated into SAGA and NuA4 in a process that requires Hsp90 and its co-chaperone, the Triple-T complex [TTT complex (Genereaux et al. 2012; Elías-Villalobos et al. 2019a)]. Tra1 contains four domains: an N-terminal “Huntington, EF3A, ATM, TOR” (HEAT) region, followed by “FRAP-ATM-TRRAP” (FAT), phosphatidylinositol 3-kinase-like (PI3K), and FAT-C-terminal (FATC) domains (Lempiäinen and Halazonetis 2009; Knutson and Hahn 2011; Baretić et al. 2017; Díaz-Santín et al. 2017). The PI3K domain is essential for Tra1 function (Mutiu et al. 2007; Knutson and Hahn 2011), yet unlike other PIKKs (Lovejoy and Cortez 2009), Tra1 lacks kinase activity (McMahon et al. 1998; Saleh et al. 1998). We recently identified three arginine residues proximal to what is the ATP-binding cleft in the PI3K domain of other PIKK proteins that, when mutated to glutamine, reduce growth, impair assembly of SAGA and NuA4 complex, and cause dysregulated transcription; we termed this allele tra1_Q3 (Berg et al. 2018). In the model yeast Saccharomyces cerevisiae, the tra1_Q3 allele results in sensitivity to high temperature, inositol auxotrophy, and decreased resistance to cell wall perturbations (Berg et al. 2018). The arginine residues are conserved among Tra1 orthologues, including in other yeast species such as C. albicans (Berg et al. 2018).

Given the connection between transcriptional regulation and stress response, it follows that chromatin-modifying factors would have important functions in mediating fungal pathogenesis, including their virulence and susceptibility to antifungal drugs (O’Kane et al. 2020). Indeed, chromatin-modifying factors including lysine (K) acetyl-transferases (KATs) such as Gcn5, Hat1, and Rtt109, and lysine deacetylases (KDACs) such as Set3 and Rpd31 are implicated in Candida virulence processes including filamentous growth and biofilm formation (da Rosa et al. 2010; Hnisz et al. 2010, 2012; Zacchi et al. 2010; Wang et al. 2013; Nobile et al. 2014; Tscherner et al. 2015; Shivarathri et al. 2019), as well as in resistance and susceptibility to antifungal drugs (Smith and Edlind 2002; Sellam et al. 2009; Robbins et al. 2012; Nobile et al. 2014; Ramírez-Zavala et al. 2014; Yu et al. 2018). The requirement of Tra1 for resistance to cell wall perturbations in S. cerevisiae (Berg et al. 2018), along with its role in both SAGA and NuA4 complexes, positions Tra1 as a potentially unique target for antifungal therapy. Despite this, the ortholog of TRA1 in the fungal pathogen C. albicans remains to be characterized and its role in antifungal resistance and fungal virulence has not been explored.

Here, we identify a key role for Tra1 in mediating resistance to antifungal drugs, as well as fungal virulence. We demonstrate that the tra1_Q3 allele increases sensitivity to the antifungal echinocandin caspofungin in S. cerevisiae. Using CRISPR editing to generate an orthologous tra1_Q3 mutant in C. albicans, we show that diminishing Tra1 increases sensitivity to caspofungin and other cell wall stressors in this fungal pathogen. Transcriptome profiling in C. albicans revealed that Tra1 is required for the transcriptional changes that occur in response to caspofungin treatment. Many of these genes were also involved in C. albicans morphogenesis and pathogenesis. Together, this work positions Tra1 as a key regulator of C. albicans drug resistance and pathogenesis, and highlights its potential as a target for antifungal therapeutics.

Materials and methods

Yeast maintenance, media, strains

Strains and plasmids used in this study are listed in Supplementary Table S1, and plasmids are listed in Supplementary Table S1. Fungal strains [S. cerevisiae (S288c derivatives) and C. albicans] were cultured on YPD (2% Bacto peptone, 1% yeast extract, and 2% glucose) or Synthetic Complete medium (with appropriate amino acids).

CRISPR design

For CRISPR-based tools used for C. albicans, including generating the tra1_Q3 mutation, and TRA1 deletion, guide RNA N20 sequences were designed based on the efficiency score and specificity using the sgRNA design tool Eukaryotic Pathogen CRISPR gRNA Design Tool (EuPaGDT) (Peng and Tarleton 2015) available at http://grna.ctegd.uga.edu. C. albicans genetic sequences were obtained from the Candida Genome Database (CGD; http://www.candidagenome.org) (Skrzypek et al. 2017).

Generation of CRISPR plasmids

CRISPR plasmids for C. albicans were generated as previously described (Shapiro et al. 2018; Halder et al. 2019; Wensing et al. 2019). Gene blocks were then cloned into the pRS252 backbone using Gibson assembly (Gibson et al. 2009). The pRS252 plasmid was digested with NgoMIV. This digested plasmid was then combined with nuclease-free water, the synthesized gene block fragment (diluted to a concentration of 50 ng/µl), and NEBuilder 2x Hifi DNA Master Mix (New England Biolabs, NEB), for a total volume of 6 μl, and incubated at 50°C for 4 h. The assembled plasmid was then transformed into chemically competent Escherichia coli DH5α using the NEB high-efficiency transformation protocol. CRISPR mutation plasmids targeting TRA1 for Q3 mutation, or knockout plasmids targeting TRA1 were engineered in pRS252 [C. albicans-optimized Cas9 plasmid (Shapiro et al. 2018; Halder et al. 2019)]. For each construct, two independent sgRNAs were synthesized: for mutation, these sgRNAs flanked the regions containing the three sites being targeted for mutagenesis; for TRA1 deletion, these sgRNAs flanked the open reading frame (ORF) of the gene. For each plasmid, a gene block (Integrated DNA Technologies, IDT) was synthesized, containing the two sgRNAs, the SNR52 sgRNA promoter, and regions of homology for Gibson assembly. The TRA1-targeting gene block additionally contained regions of homology upstream and downstream of the TRA1 ORF.

C. albicans transformation

C. albicans cells were transformed as previously described (Halder et al. 2019). Briefly, the plasmids to be transformed were miniprepped and plasmid quality and purity were determined using a Nanodrop spectrophotometer (Tecan). Cas9 plasmids were linearized with the PacI restriction enzyme (NEB) to enable plasmid integration at the NEUT5L locus. CRISPR plasmids were co-transformed with repair templates (gRS11 for tra1_Q3 containing Q3 mutations, gRS83 for tra1Δ). Linearized plasmids, repair templates, and C. albicans cells were incubated with 800 µl 50% polyethylene glycol, 100 µl 10x Tris-EDTA buffer, 100 µl 1 M lithium acetate (pH 7.4), 40 µl salmon sperm DNA, and 20 µl 2 M dithiothreitol, and incubated at 30°C for 1 h and at 42°C for 45 min. Post-transformation, cells were grown in YPD medium for 4 h at 30°C with shaking and transformants were selected for on YPD agar plates containing 200 µg/ml nourseothricin (Jena Biosciences). C. albicans transformed colonies were PCR-verified for presence of the Cas9 construct and/or specific gene deletions. The presence of the three targeted point mutations (R3471Q, R3472Q, and R3538Q) in TRA1 was confirmed through Sanger sequencing.

CFU spot platings

Wild-type and tra1_Q3 strains were grown in 5 ml YPD overnight at 30°C. Strains were then diluted to an OD₆₀₀ of 0.05 and grown to early log phase, with an OD₆₀₀ of 0.2, by shaking at 30°C for 2–3 h. Since tra1Δ mutants were slow-growing, overnight cultures for these strains, in addition to their respective wild-type strains, were not subcultured to an OD₆₀₀ of 0.05, but rather diluted directly to an OD₆₀₀ of 0.2. Overnight cultures of tra1Δ mutants (transformant 1 and 2) and the wild-type strains were photographed after 15 h of growth. One hundred microliters of each strain was transferred to the first well of each row of a 96-well plate and 10-fold serial dilutions made in YPD. Five microliters of each row was spotted onto plain YPD plates, or YPD plates containing 25 μg/ml calcofluor white or 0.25 μg/ml caspofungin. Each plate was set up in triplicate. Plates were incubated at 30°C for 2 days.

RNA preparation and sequencing

For C. albicans, RNA samples were generated in triplicate for each mutant. The following triplicate sets were grown overnight in 50 ml inoculated YPD tubes, for a total of 12 tubes: wild type without drug, wild type grown with 10 ng/ml caspofungin, tra1_Q3 mutant without drug, and tra1_Q3 mutant grown with 10 ng/ml caspofungin. The appropriate samples were inoculated with drug when at an OD₆₀₀ of 0.1, and then grown to an OD₆₀₀ of 0.2 with drug for approximately 2 h at 30°C. RNA samples were then extracted using the Presto Mini RNA Yeast Kit (Geneaid). For S. cerevisiae, RNA was extracted from three replicates from each strain at early log phase growth in synthetic complete medium. RNA was extracted using the MasterPure Yeast RNA purification kit (Lucigen) according to the manufacturer’s instructions. Sample purity was checked using a tapestation bioanalyzer to ensure a RIN value of 8.0 or higher before being vacuum dried. Dried sample pellets were sent to Genewiz for further analysis.

Total RNA sequencing was performed by Genewiz (South Plainfield, NJ, USA). Stranded Illumina TruSeq cDNA libraries with poly dT enrichment were prepared from high-quality total RNA (RIN > 8). Libraries were sequenced on an Illumina HiSeq, yielding between 26.4 and 37.9 million 150 bp paired-end sequencing reads per sample. The data have been deposited in NCBI’s Gene Expression Omnibus (Edgar et al. 2002) and are accessible through GEO Series accession number GSE168955 for S. cerevisiae (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE168955) and GSE168988 for C. albicans (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE168988).

Quality control, trimming, read alignment, and differential gene expression analysis

FASTQ files were analyzed using a customized bioinformatics workflow. Adapter sequences and low-quality bases were trimmed using the default settings of Trimmomatic (Bolger et al. 2014). Sequence quality was analyzed using FastQC (http://www.bioinformatics.babraham.ac.uk/projects/fastqc/). Sequence reads were aligned to the S. cerevisiae S288C reference genome (assembly R64-2-1; https://www.yeastgenome.org/) or the C. albicans SC5314 reference genome (assembly 21; http://www.candidagenome.org) using STAR (Dobin et al. 2013). Only reads that were uniquely mapped to the reference genome were kept. Read counts for each gene were summarized using featureCounts (Liao et al. 2014). Differential expression analysis was performed using the DESeq2 R package (Love et al. 2014) using a custom R script (Supplementary File S5) with a Benjamini-Hochberg adjusted P-value cut-off of ≤0.05. The volcano plot presented in Supplementary Figure S3 was generated using VolcaNoseR (Goedhart and Luijsterburg 2020).

Filamentation assay and microscopy

Wild-type and mutant strains were grown overnight in 5 ml of YPD at 30°C. OD₆₀₀ reads were taken and strain growth was normalized by diluting samples to OD₆₀₀ = 0.2. Cells were subcultured and incubated in YPD with 10% serum or in Spider medium to induce filamentation. Strains induced with serum or grown in Spider medium were incubated at 37°C, while strains grown in nonfilamentous conditions were incubated at 30°C, for four hours. Filamentation was observed via bright-field microscopy.

Growth curves

Overnight cultures of 5 ml YPD inoculated with C. albicans were grown at 30°C, with shaking at 250 rpm. Growth of the strains was evaluated by measuring optical density at 600 nm and strain growth was normalized by diluting to OD₆₀₀ = 0.1. In a 96-well plate containing 100 μl of fresh YPD, 100 μl of each sample was added, followed by the addition of 50 μl of mineral oil to prevent evaporation. OD₆₀₀ was determined every 15 min for 24 h in a microplate reader, with shaking between reads. The data were graphed using Graphpad Prism version 8.

Biofilm growth

Wild-type and tra1_Q3 strains were incubated overnight in 5 ml YPD at 37°C and 250 rpm. The cultures were normalized to the same OD₆₀₀ by diluting with RPMI liquid medium. One hundred microliters of each subcultured strain was distributed into flat bottom 96-well polystyrene plates containing 100 μl of RPMI in each well. A row containing 200 μl of RPMI was used as a negative control. The 96-well plates were wrapped in aluminum foil, then incubated at 37°C for 72 h. One hundred and twenty microliters of planktonic cells was removed from each well, dispensed into new 96-well plates, and OD₆₀₀ determined in a microplate reader. The biofilm plates were washed twice with 200 μl of 1x PBS buffer and dried in a fume hood for ∼1 h. Next, 90 μl of XTT sodium salt (1 mg/ml) and 10 μl of phenazine methosulfate (PMS; 0.32 mg/ml) were added sequentially to every well, the plates were wrapped in aluminum foil, and incubated for 2 h at 30°C. OD₄₉₀ was determined in a microplate reader and the XTT biofilm growth values normalized to planktonic cell growth.

Macrophage cell lines

Immortalized macrophages, of BALB/c murine origin, were used to conduct macrophage infection assays. Prior to infection, cells were passaged into three 150 mm tissue culture dishes of fresh Dulbecco′s Modified Eagle′s Medium (DMEM; Gibco) medium, supplemented with 2 mM L-glutamine (Gibco), 2 mM penicillin-streptomycin (Lonza), and 10% heat-inactivated fetal bovine serum (Life Technologies). These cells were then incubated at 37°C and 5% CO₂, for three days to grow up to 70–80% confluence. On infection day, cells were washed and collected in DMEM medium lacking penicillin-streptomycin (- pen - strep). The concentration of living cells was determined using the Countess™ II automated cell counter (Invitrogen). The appropriate volume of cells was added to each well of a 12-well tissue culture plate, to obtain 500,000 cells in 1 ml medium. Cells were incubated for 3 h at 37°C and 5% CO₂, to allow adherence to the plate surface prior to infection.

C. albicans-macrophage infection

Overnight cultures of C. albicans strains were grown in 5 ml YPD at 37°C and 250 rpm, the day before infection. Cells were then subcultured in 5 ml of fresh YPD and grown to an OD₆₀₀ of 1.0. Cells were washed and resuspended in 1 ml DMEM (- pen - strep). Cell count was determined on a hemocytometer under a light microscope. C. albicans strains were added to macrophages at a ratio of 1 yeast cell: 10 macrophages. Assuming 500,000 macrophages are in each well of the tissue culture plate, the appropriate volume of C. albicans cells was added to obtain 50,000 cells/well. DMEM (- pen - strep) medium, as well as uninfected macrophages, were included as controls.

C. albicans and macrophages were co-incubated for 1 h at 37°C and 5% CO₂ to allow infection. All further incubations were at 37°C and 5% CO₂. Wells were washed with 1x sterile PBS (without calcium or magnesium) and cells incubated 30 min with DMEM (- pen - strep), supplemented with 5 µg/ml caspofungin to kill residual C. albicans. Wells were washed three times with 1x PBS and provided with 1 ml of DMEM (- pen - strep) medium. Plates were incubated and the cytotoxicity of each C. albicans strain was measured using the LDH assay at the 1-, 3-, 6-, and 18-h time points. The LDH assay was conducted using the CytoTox 96^® NonRadioactive Cytotoxicity Assay kit (Promega). At each time point, the supernatant from each well was collected and 1 ml of 1.2% Triton X (diluted with PBS) was added to lyse cells for 30 min in the dark. On a 96-well plate, 50 µl of LDH substrate was added to 50 µl of both the supernatant and cell lysate for each treatment condition, as well as the untreated and media control conditions. Plates were incubated in the dark for 30 min, after which 50 µl of the stop solution was added. The OD₄₉₀ was measured for each condition, and the % cytotoxicity of each strain (treatment) was calculated using these OD₄₉₀ values in the following equation: % cytotoxicity = (treatment - media control)/(uninfected control - media control) × 100%.

C. elegans-C. albicans killing assay

A C. elegans-based analysis of fungal virulence was performed, as previously described (Kim et al. 2020; Rosiana et al. 2021). Briefly, C. elegans glp-4 worms were bleached and resuspended to extract their eggs and establish a fresh stock of worms. Approximately 5000–6000 worms were dispensed onto 3–4 superfood agar plates, which were then left overnight at room temperature and then moved to a 25°C static incubator for the next two nights. Cultures of C. albicans strains were made using 5 ml of YPD and left to incubate at 30°C and 250 rpm overnight. The following day, 10 μl of the culture was spread onto BHI-kanamycin (Kan) plates, which were incubated overnight at 30°C static. The C. elegans superfood agar plates were washed with ∼6 ml of S Basal to dislodge the worms and then the worms were washed 3 times with S Basal. A solution of 2 worms per μl S Basal was made and 80–100 worms were dispensed onto each C. albicans-inoculated BHI-Kan plate. The plates were then stored in a 25°C incubator. The worms were scored over a period of 7 days by counting and removing dead worms with a platinum wire worm pick under a stereomicroscope. A Kaplan-Meier survival graph was generated using Graphpad Prism.

Results

The tra1_Q3 allele increases susceptibility to caspofungin in S. cerevisiae and C. albicans

Amongst Tra1 orthologues, the positively charged residues at positions 3389, 3390, and 3456 are highly conserved (Berg et al. 2018). When mapped onto the cryo-EM structure of Tra1 (Díaz-Santín et al. 2017), these conserved arginine residues fall within or near the ATP-binding cleft (Berg et al. 2018). Although the PI3K domain of Tra1 lacks key residues required for ATP binding, altering these three arginine residues to glutamine, an allele termed tra1_Q3, reduces assembly of Tra1 into SAGA and NuA4 complexes in S. cerevisiae, demonstrating the importance of the cleft (Berg et al. 2018). To further assess the importance of the Tra1 PI3K domain on SAGA/NuA4-mediated transcription, we performed RNA sequencing (RNA-seq) to identify genes and pathways that are differentially expressed in the S. cerevisiae tra1_Q3 strain compared to a wild-type strain. We identified 1724 and 1792 genes that were significantly up- and down-regulated, respectively (adjusted P-value < 0.05; Figure 1A and Supplementary File S1). In the set of genes upregulated greater than twofold in tra1_Q3 relative to wild-type, GO term analysis identified enrichment for genes involved in cell wall organization (Figure 1B). Down-regulated genes with greater than 2-fold change were enriched for roles in polyphosphate metabolism (Figure 1C).

Figure 1

Transcriptome analysis of S. cerevisiae expressing tra1_Q3. (A) Volcano plot of changes in gene expression as determined by RNA-seq in strains expressing tra1_Q3 relative to a strain expressing wild-type TRA1. Differentially expressed genes with an adjusted P-value <0.05 are colored light blue (up regulated in tra1_Q3 relative to TRA1) and dark blue (down regulated in tra1_Q3 relative to TRA1). (B) Significantly enriched GO biological processes were determined from the set of significant genes with >2-fold up regulation (adjusted P < 0.05) in tra1_Q3 relative to TRA1. (C) Significantly enriched GO biological processes were determined from the set of genes with >2-fold down regulation (adjusted P < 0.05) in tra1_Q3 relative to TRA1. (D) Differentially expressed genes in tra1_Q3 were analyzed for transcription factor associations using YEASTRACT+. GO analysis was performed with the percentage of identified transcription factors (162) for each GO term shown in the bar graph. (E) Rlm1 associations with differentially expressed genes in the tra1_Q3 strain. The experimental evidence underlying each regulatory association (solid lines for DNA-binding evidence; dashed lines for expression evidence), as well as the sign of the interaction—positive (green), negative (red), positive and negative (brown), or undefined (black) are displayed.

We also identified differential expression of genes involved in various stress responses in the tra1_Q3 strain, in agreement with previous studies demonstrating that SAGA and NuA4 regulate stress responsive genes (Huisinga and Pugh 2004; Mitchell et al. 2008; Helmlinger et al. 2011). These included genes involved in the cell wall integrity pathway (Levin 2011) (PKH1, MTL1, KSS1, SWI6, MSG5, WSC3, PTP2, and PTP3 were downregulated and SAC7 and RHO1 were upregulated). Compromised cell wall integrity pathway (CWI) signaling leads to cell wall remodeling, confirmed by enrichment of genes associated with cell wall organization among upregulated genes in our dataset. Indeed, we observed upregulation of genes encoding proteins involved in cell wall architecture such as PIR3, PST1, YPG1, and SPI1 that have been shown to be increased in cell wall mutants (Lagorce et al. 2003). Previous work also demonstrated a role for Tra1 in resistance to cell wall stress (Mutiu et al. 2007; Hoke et al. 2010; Sanz et al. 2016; Berg et al. 2018). We then analyzed differentially regulated genes (392 genes, log₂ fold change > 1, P < 0.05) for known association with transcription factors using yeastract (www.yeastract.org) (Monteiro et al. 2020) (Figure 1D, Supplementary File S2). GO analysis of identified transcription factors revealed factors linked to processes associated with PolII transcription, in agreement with the broad role of SAGA in global transcriptional regulation (Baptista et al. 2018). Other transcription factor associations (Hsf1 and Hac1) highlight the role of Tra1 in stress responses. Interestingly, our analysis also shows significant enrichment in Rlm1 (Figure 1E) target genes and Sfp1 targets (Supplementary Figure S1). Sfp1 interacts with Tra1 (Lempiäinen et al. 2009) and we recently demonstrated that Sfp1 regulates Tra1/SAGA-driven gene expression in response to protein misfolding (Jiang et al. 2019). Rlm1 controls gene expression of factors involved in CWI (Levin 2011), consistent with the tra1_Q3 causing increased sensitivity to cell wall perturbation (Berg et al. 2018). Moreover, SAGA recruitment to the promoters of CWI target genes requires Rlm1 (Sanz et al. 2016).

As many antifungals target the cell wall or plasma membrane and induce stress responses (Shapiro et al. 2011), we hypothesized that Tra1 might regulate the antifungal response. We therefore assessed whether the tra1_Q3 strain was sensitive to the antifungal agent caspofungin, an echinocandin that causes cell wall stress. Growth of the tra1_Q3 strain was significantly reduced by caspofungin in both solid and liquid media conditions (Figure 2, A and B).

Figure 2

S. cerevisiae TRA1 deficient strains are hypersensitive to antifungal stress. (A) Growth of the S. cerevisiae tra1_Q3 strain on solid medium is impaired by 0.25 µg/ml caspofungin compared to a wild-type strain. S. cerevisiae strains were serially diluted (1:10) and spotted onto solid YPD agar media with or without 0.25 µg/ml caspofungin. (B) Growth of the S. cerevisiae tra1_Q3 strain is hypersensitive to 0.25 µg/ml caspofungin in liquid medium. S. cerevisiae strains were grown in liquid YPD medium with or without 0.25 µg/ml caspofungin continuously over 1000 min. Growth was determined by optical density (OD₆₀₀).

Hypothesizing that Tra1 could be a potential target for antifungal therapy, we next examined whether Tra1 was similarly involved in sensitivity to cell wall stressors in the fungal pathogen C. albicans. First, we identified the three conserved positively charged residues (K3471, R3472, and R3538) that correspond to the residues mutated in the S. cerevisiae tra1_Q3 (Figure 3A). We used CRISPR-Cas9-based genome editing tools (Halder et al. 2019) to modify these residues to glutamine for both alleles of TRA1 in diploid C. albicans. We generated two independent tra1_Q3 strains in C. albicans and assessed the sensitivity of these strains to cell wall stressors, as compared with a wild-type strain. As was the case in S. cerevisiae, tra1_Q3 rendered C. albicans significantly more sensitive to the presence of the cell wall stressor calcofluor white, as well as the cell wall-targeting antifungal caspofungin (Figure 3B; ANOVA P < 0.05).

Figure 3

C. albicans tra1_Q3 strains are hypersensitive to cell wall stressors. (A) Alignment of TRA1 from S. cerevisiae and C. albicans, highlighting (red asterisks) the conserved positively charged residues (K3471, R3472, and R3538) proximal to the PI3K ATP-binding cleft, which correspond to residues mutated in the S. cerevisiae tra1_Q3 strain. (B) tra1_Q3 results in hypersensitivity caspofungin or calcofluor white in solid medium. C. albicans strains were serially diluted (1:10 dilutions) and spotted onto solid YPD agar media with or without 0.25 µg/ml caspofungin or 25 µg/ml calcofluor white. CFU/ml counts for three or four replicate plates per condition were quantified, and tra1_Q3 strains CFU/ml on caspofungin or calcofluor white were significantly different from wild type, ANOVA P < 0.05.

Tra1 is essential for viability in S. cerevisiae and other yeasts, with the exception of the fission yeast Schizosaccharomyces pombe, where two paralogs exist; SpTra1 specifically associates with SAGA and is not essential, while SpTra2 associates with NuA4 and is essential (Helmlinger et al. 2011). Previous reports have unexpectedly suggested that Tra1 may not be essential in C. albicans (O’Meara et al. 2015; Segal et al. 2018). Therefore, to investigate the requirement for TRA1 for C. albicans viability, we attempted to generate a TRA1 knockout strain using a CRISPR approach. While we succeeded in knocking out the gene (Supplementary Figure S2A), the resulting strain displayed a severe growth defect (Supplementary Figure S2, B and C) suggesting that TRA1, while not strictly essential, is required for proper growth. Based on the significant fitness defect associated with TRA1 deletion, we focus on the tra1_Q3 mutant strains for subsequent analysis.

RNA-seq analysis reveals the transcriptional signature of tra1_Q3 in response to caspofungin

Given the role of Tra1 as an essential component of SAGA and NuA4 histone acetyltransferase complexes, and its role in sensitivity to the antifungal caspofungin, we next sought to explore the role of TRA1 in mediating the fungal transcriptional response to treatment with caspofungin. Wild-type and tra1_Q3 strains of C. albicans were grown in the presence or absence of caspofungin, and RNA-seq was performed. Overall, ∼45% of the variation in the resultant transcriptomic dataset was explained by treatment with caspofungin, and ∼20% of the variation explained by the tra1_Q3 mutation (Figure 4A). For the untreated tra1_Q3 strain, 1514 genes were differentially expressed, 841 downregulated and 673 upregulated when compared to wild-type (adjusted P-value < 0.05). Caspofungin treatment in wild-type cells resulted in 3113 genes being differentially expressed, 1723 downregulated and 1390 upregulated. Compared to the wild-type C. albicans strain, the number of genes differentially regulated upon caspofungin treatment in the tra1_Q3 expressing strain was reduced to 1763, 766 downregulated and 997 upregulated. A comprehensive list of significant genes and the fold-change expression data for each comparison can be found in Supplementary File S3.

Figure 4

Transcriptome analysis of C. albicans expressing tra1_Q3 in response to caspofungin. (A) Principal component analysis of centered log ratio normalized reads from C. albicans expressing either wild type TRA1 or tra1_Q3 with and without caspofungin treatment. Each point represents one biological replicate (n = 3). (B) Volcano plot of genes that respond differently to caspofungin treatment in the strain expressing tra1_Q3 relative to the wild-type strain. Genes with log₂ fold change response > 1 (adjusted P-value < 0.05) are colored red and genes with log₂ fold change response <−1 (adjusted P-value < 0.05) are colored dark red. (C) Heatmap of hierarchical clustered genes with different responses to caspofungin in tra1_Q3 expressing strains relative to wild type strains (log₂ fold change > |1|, adjusted P-value < 0.05). Fold-change for each gene is the average from three biological replicates. Genes that are upregulated and downregulated upon caspofungin treatment are colored yellow and blue, respectively.

From this dataset, we identified the genes responding most differently to treatment with caspofungin in the presence of tra1_Q3, as compared to caspofungin treatment of the wild-type strain and thus possibly underpinning the genotype-condition interaction (Figure 4B). There were 68 genes that had a significantly different response based on this genotype-condition interaction (log₂ fold change > 1, adjusted P-value < 0.05). A heat map representing the changes in expression upon caspofungin treatment for wild type and tra1_Q3 strains is shown in Figure 4C. The most pronounced differences were seen for HPD1, a dehydrogenase involved in the degradation of toxic propionyl-CoA, and PHR1, a cell surface glycosidase. Although there was no significant gene ontology (GO) term enrichment in this smaller subset of differentially regulated genes, the genes annotated with a known biological process included many involved in cellular transport (ADH3, EVP1, FTH1, HAK1, HOF1, HPD1, MEP1, MOB1, NGT1, OPT7, PHHB, QDR1, and TRY6), filamentous growth (ASH1, ERG1, ERG3, PHHB, PHR1, and WOR2), biofilm growth (PHR1, QDR1, and TRY6), and response to chemicals, drugs, and stress (ASH1, ERG1, ERG3, HOF1, MEP1, MMS21, PHHB, HAK1, PHR1, and QDR1).

The genes uniquely regulated by TRA1 upon caspofungin treatment were then analyzed for regulatory associations (including documented DNA-binding and expression-based evidence) with known transcription factors using the PathoYeastract portal (Monteiro et al. 2020). We identified several transcription factors including Pho4 (known for being regulated by SAGA/NuA4 in S. cerevisiae  Barbaric et al. 2003; Nourani et al. 2004; Adkins et al. 2007) as well as Nrg1 and Ron1, with known roles in filamentous growth, biofilm formation and response to stress and hyphal growth (Figure 5A and Supplementary File S4) (Murad 2001; Moran et al. 2007; Cleary and Saville 2010; Urrialde et al. 2016; Song et al. 2020). Regulatory associations of Pho4, Nrg1, and Ron1 with genes differentially expressed in tra1_Q3 are shown in Figure 5B. Other transcription factors previously linked to SAGA/NuA4 include Snf5, Skn7, Sko1, Rim101, Fkh2, and Mcm1 (Proft and Struhl 2002; Chandy et al. 2006; Sellam et al. 2009; Lin et al. 2012; Dewhurst-Maridor et al. 2017). Among identified factors, Sko1 and Cas5, regulate the response to the caspofungin-induced cell wall damage in C. albicans (Bruno et al. 2006; Heredia et al. 2020a,b). Taken together, these data support an important role for Tra1 in regulating the transcriptional response to caspofungin.

Figure 5

Transcription factor association underpinning the tra1_Q3 response to caspofungin in C. albicans. (A) Differentially expressed genes in tra1_Q3 in response to caspofungin treatment were analysed for transcription factor associations using YEASTRACT+. GO analysis was performed and the percentage of identified transcription factors (73) for each GO term shown in the bar graph. (B) Pho4, Nrg1, and Ron1 associations with differentially expressed genes in the tra1_Q3 strain in response to caspofungin treatment. The experimental evidence underlying each regulatory association (full lines for DNA-binding evidence; dashed for expression evidence), as well as the sign of the interaction—positive (green), negative (red), positive and negative (brown), or undefined (black) are displayed.

TRA1 plays a role in C. albicans pathogenesis

Since many differentially regulated genes unique to the tra1_Q3 response to caspofungin had annotated roles in fungal pathogenesis processes, we explored whether TRA1 is involved in pathogenesis phenotypes in C. albicans. One key pathogenesis-associated phenotype for C. albicans is the ability to undergo a reversible morphological transition between yeast and filamentous growth (Shapiro et al. 2011). We monitored wild-type and tra1_Q3 strains of C. albicans for their ability to undergo cellular morphogenesis and filamentation in response to diverse media conditions known to induce filamentation, including growth in medium containing 10% serum and Spider medium. We found that filamentation of the tra1_Q3 strain isolates was impaired compared to the wild-type strain (Figure 6A), suggesting that TRA1 is required for the filamentous growth transition in C. albicans. This filamentous growth defect is specific to filamentation, and is not an overall growth defect of this mutant (Supplementary Figure S2D). Given the important role that morphological transitions play in biofilm formation in C. albicans (Lohse et al. 2018), we further investigated the role of TRA1 in biofilm formation. We allowed C. albicans biofilms to form in multi-well plates, removed planktonic cells, and quantified the presence of metabolically active adherent fungal cells as a measure of biofilm growth. The tra1_Q3 isolates were less capable of forming robust biofilms, compared to a wild-type strain of C. albicans (Figure 6B;  P < 0.0001, ANOVA).

Figure 6

C. albicans tra1_Q3 impairs pathogenesis-associated traits. (A) C. albicans tra1_Q3 mutants are impaired in the morphogenetic transition from yeast to filamentous growth in YPD media containing 10% serum or in Spider media. C. albicans strains were grown in YPD media alone at 30°C, or in YPD containing 10% serum or in Spider media at 37°C for 4 h. Cells were observed by bright-field microscopy. Scale bar = 50 µm (B) C. albicans tra1_Q3 mutants are impaired in biofilm formation relative to a wild-type strain (ANOVA, P < 0.0001 ****). Biofilm growth was quantified by an XTT metabolic readout, measured at OD₄₉₀ and normalized to planktonic growth. Each point represents an independent biofilm replicate. (C) The C. albicans tra1_Q3 strain has reduced macrophage cytotoxicity relative to a wild-type strain (ANOVA, P < 0.0001 ****, P < 0.001 ***, error bars represent standard deviation). Macrophage cytotoxicity was quantified by measuring LDH after 6 h of co-incubation with C. albicans strains. For LDH measurement, OD₄₉₀ was measured, and the percent cytotoxicity of each strain calculated using these OD₄₉₀ values relative to OD₄₉₀ of uninfected control macrophages. Each point represents a replicate co-incubation assay. (D) The C. albicans tra1_Q3 strain has reduced virulence relative to a wild-type strain, using a C. elegans model of fungal infection. C. elegans survival was monitored over 7 days, and survival was plotted with a Kaplan-Meier survival curve.

The importance of TRA1 in mediating filamentation and in biofilm formation led us to determine whether it was also involved in the interaction of C. albicans with mammalian host cells, as C. albicans morphogenesis is linked with host cell interaction and escape (Gow et al. 2011). We performed a macrophage-C. albicans infection assay, where fungal cells were co-incubated with BALB/c immortalized murine macrophage cells. We monitored lactate dehydrogenase (LDH) release as a measure of macrophage cell death upon infection by wild-type or tra1_Q3 C. albicans strains, relative to uninfected macrophages. Macrophage cells infected with tra1_Q3 strains generated significantly less LDH compared to macrophages infected with wild-type C. albicans (Figure 6C;  P < 0.001, ANOVA), indicating less host cytotoxicity associated with tra1_Q3, and suggesting a role for TRA1 in the infection and killing of host macrophage cells.

Finally, to assess the role of TRA1 in fungal pathogenesis, we exploited a C. elegans infection model to monitor virulence of the tra1_Q3 mutant relative to a wild-type strain. We found that over the seven-day time course, the tra1_Q3 mutant had reduced virulence compared to the wild-type strain (Figure 6D), suggesting an important role for TRA1 in mediating fungal virulence.

Discussion

We previously demonstrated that S. cerevisiae Tra1_Q3 is poorly integrated into the SAGA and NuA4 complexes, compromises Tra1-dependent transcription, and mislocalizes to the cytoplasm (Berg et al. 2018). The transcriptome analysis presented here shows that Tra1 affects multiple cellular processes, in agreement with the role of SAGA as a general cofactor for RNA polymerase II transcription (Bonnet et al. 2014; Baptista et al. 2018; Berg et al. 2018; Bruzzone et al. 2018). Differential expression of cell wall genes, regulatory association with transcription factors such as Rlm1 and Hsf1, and sensitivity to caspofungin are consistent with the role of Tra1 in stress responses (Huisinga and Pugh 2004), and are in agreement with our previous analysis of other Tra1 derivatives (Mutiu et al. 2007; Hoke et al. 2008). The tra1_Q3 mutant displayed reduced expression of genes associated with SAGA-regulated transcription such as genes involved in phosphate homeostasis. Regulation of proper intracellular phosphate levels is crucial for cellular adaptation to multiple stresses (Austin and Mayer 2020). Interestingly, components of the phosphate transport and metabolism pathways control antifungal resistance and pathogenicity of fungal pathogens (Liu et al. 2017, 2018; Köhler et al. 2020; Peng et al. 2020).

This work is the first to characterize the role of TRA1 in the fungal pathogen C. albicans. We found that, similar to S. cerevisiae, the C. albicans tra1_Q3 strain is highly sensitive to cell wall-targeting stressors, including calcofluor white and the antifungal drug caspofungin. Of note, the tra1_Q3 allele impairs several measures of fungal pathogenesis, including filamentation, biofilm formation, macrophage cytotoxicity, and virulence in a C. elegans model of fungal infection. Many of these pathogenesis traits are linked, as filamentation is a critical part of biofilm formation and interaction with host immune cells (Whiteway and Bachewich 2007; Gow et al. 2011; Shapiro et al. 2011). These findings add to a growing literature highlighting the key role of histone acetyltransferases and histone deacetylases in mediating fungal virulence (da Rosa et al. 2010; Hnisz et al. 2010, 2012; Zacchi et al. 2010; Wang et al. 2013; Nobile et al. 2014; Tscherner et al. 2015; Shivarathri et al. 2019) and resistance to antifungal drugs (Smith and Edlind 2002; Sellam et al. 2009; Robbins et al. 2012; Nobile et al. 2014; Ramírez-Zavala et al. 2014; Yu et al. 2018; Shivarathri et al. 2019). Recent work with the histone acetyltransferase Gcn5 in C. albicans has focused on unraveling the mechanisms by which histone-modifying enzymes alter fungal virulence and sensitivity to antifungal agents, including caspofungin (Shivarathri et al. 2019). Similar to TRA1, loss of GCN5 in C. albicans is associated with defects in filamentation, hypersensitivity to caspofungin, and other virulence and stress-related phenotypes (Shivarathri et al. 2019). GCN5-related caspofungin susceptibility may be linked to Gcn5-mediated regulation of the master transcriptional regulator Efg1 (Shivarathri et al. 2019), which is involved in caspofungin susceptibility (Gregori et al. 2011). Loss of Gcn5 is associated with altered MAPK signaling, which is involved in remodeling of the fungal cell wall, as well as filamentation and other virulence processes (Monge et al. 2006; Román et al. 2007; Chow et al. 2018; Shivarathri et al. 2019). Interestingly, Efg1 is one of the transcription factors associated with the genes underpinning the Tra1-regulated caspofungin response (Supplementary File S4). Despite the functional similarities, we find little overlap between our tra1_Q3 dataset and the transcriptional profile of C. albicans cells depleted of GCN5 (Supplementary Figure S3). It is difficult to evaluate if these differences emerge from the comparison of a loss-of-function Tra1 mutant with a complete GCN5 knockout, the role of Tra1 in both SAGA and NuA4, or the possibility that C. albicans Gcn5 regulates a subset of genes in the absence of a functional Tra1, as it was recently suggested for S. cerevisiae (Bruzzone et al. 2018).

Another interesting result is the absence of TRA1 upregulation in the C. albicans tra1_Q3 strain, which contrasts to what we observed in S. cerevisiae (3.2-fold upregulated) (Berg et al. 2018). Moreover, we found only four genes (HSP30, IZH1, OPT1, and RDS3) that were differentially regulated by Tra1_Q3 in both species. Thus, it appears that several genes, including TRA1, have undergone a transcriptional rewiring between S. cerevisiae and C. albicans in response to loss of Tra1 function. In contrast to the TRA1 knockout strain, the tra1_Q3 mutant displayed no growth defects in C. albicans. Our previous work in S. cerevisiae showed that while displaying impaired assembly of both SAGA and NuA4, tra1_Q3 supports viability (Berg et al. 2018). The breadth of functions regulated by Tra1 is reflected by the various phenotypes observed in mutational analysis. Tra1 appears to have both overlapping and distinct functions with SAGA and NuA4. Which of those functions are required to support viability remains unclear (Mutiu et al. 2007; Hoke et al. 2008, 2010; Genereaux et al. 2012; Lin et al. 2012). Future investigations on the role of C. albicans Tra1 in the recruitment of the acetyltransferase Gcn5 and Esa1 to promoters, as well as Tra1 regulation of histone acetylation, will help clarify these issues. Nevertheless, the key role of these enzymes in fungal virulence and drug susceptibility indicates their potential as antifungal therapeutic targets (Bauer et al. 2016; Kuchler et al. 2016).

The C. albicans tra1_Q3 strain had altered phenotypes associated with both antifungal drug resistance, as well as pathogenicity traits such as filamentation. This connection between drug resistance and pathogenesis has been well established amongst pathogenic Candida species, and numerous signaling pathways jointly mediate these two important facets of fungal biology (Sharma et al. 2019). One central regulator of fungal pathogenesis and drug resistance is the essential molecular chaperone Hsp90, which plays a key role in C. albicans morphogenesis, biofilm formation, resistance to azole and echinocandin antifungals, and virulence in animal models of infection (Cowen and Lindquist 2005; Shapiro et al. 2009; Singh et al. 2009; Robbins et al. 2011; O’Meara et al. 2017). Hsp90 itself is regulated by lysine deacetylases including Hos2, Hda1, Rpd3, and Rpd31, which modulate Hsp90’s role in antifungal drug resistance (Robbins et al. 2012) and morphogenesis (Li et al. 2017), and numerous downstream cellular pathways are involved in Hsp90-mediated regulation of virulence and drug resistance, including protein kinase A (PKA), and MAPK signaling pathways (Shapiro et al. 2009; LaFayette et al. 2010; Diezmann et al. 2012). Recent work in the model yeast S. pombe found that the Tra1 and Tra2 proteins require Hsp90 along with a cochaperone, the Triple-T complex, for integration into the SAGA and NuA4 complexes (Elías-Villalobos et al. 2019a). If the Tra1-Hsp90 interaction is conserved in C. albicans, this interaction may provide at least one rationale for how Hsp90 regulates morphogenesis and virulence.

Our results indicate that Tra1 is an attractive target for combinational antifungal therapy together with current compounds such as caspofungin. Although the exact function of Tra1’s PI3K domain is unknown, it is critical for fungal growth and survival [Supplementary Figure S2, B and C, discussed in more details in Berg et al. (2018)]. Like other PIKK family members such as mTor, the activity of the PI3K domain of Tra1 should be druggable. The putative ATP-binding cleft represents an ideal target for small molecule inhibitors and these efforts will be further facilitated by recent Tra1 structural studies (Díaz-Santín et al. 2017; Sharov et al. 2017).

Data availability

Supplemental figures and files have been uploaded through the GSA Figshare portal: https://doi.org/10.25386/genetics.15053076. The data from the RNA-seq experiments have been deposited in NCBI’s Gene Expression Omnibus (Edgar et al. 2002) and are accessible through GEO Series accession number GSE168955 for S. cerevisiae (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE168955) and GSE168988 for C. albicans (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE168988). C. albicans CRISPR plasmid backbones are available via Addgene, Addgene ID: 89576.

Funding

This work was supported by a Canadian Institutes of Health Research (CIHR) Project Grant (PJT 162195) and Natural Sciences and Engineering Research Council (NSERC) Discovery Grant (RGPIN-2018-4914) to R.S.S. P.L. is supported by an NSERC Discovery Grant (RGPIN-2015-06400) and a Canadian Foundation for Innovation (CFI) John R. Evans Leader Fund Grant (65183). C.J.B. is supported by an NSERC Discovery Grant (RGPIN-2015-04394). Y.J. was supported by a Masters to PhD Transfer Scholarship from the Dean of the Schulich Faculty of Medicine and Dentistry at Western University. M.D.B. was supported by an NSERC Alexander Graham Bell Canada Graduate Scholarship.

Conflicts of interest

The authors declare that there is no conflict of interest.

Author notes

Iqra Razzaq, Matthew D. Berg, Patrick Lajoie and Rebecca S. Shapiro contributed equally to this work.