Mitochondria Influence CDR1 Efflux Pump Activity, Hog1-Mediated Oxidative Stress Pathway, Iron Homeostasis, and Ergosterol Levels in Candida albicans

Abstract

Mitochondrial dysfunction in Candida albicans is known to be associated with drug susceptibility, cell wall integrity, phospholipid homeostasis, and virulence. In this study, we deleted CaFZO1, a key component required during biogenesis of functional mitochondria. Cells with FZO1 deleted displayed fragmented mitochondria, mitochondrial genome loss, and reduced mitochondrial membrane potential and were rendered sensitive to azoles and peroxide. In order to understand the cellular response to dysfunctional mitochondria, genome-wide expression profiling of fzo1Δ/Δ cells was performed. Our results show that the increased susceptibility to azoles was likely due to reduced efflux activity of CDR efflux pumps, caused by the missorting of Cdr1p into the vacuole. In addition, fzo1Δ/Δ cells showed upregulation of genes involved in iron assimilation, in iron-sufficient conditions, characteristic of iron-starved cells. One of the consequent effects was downregulation of genes of the ergosterol biosynthesis pathway with a commensurate decrease in cellular ergosterol levels. We therefore connect deregulated iron metabolism to ergosterol biosynthesis pathway in response to dysfunctional mitochondria. Impaired activation of the Hog1 pathway in the mutant was the basis for increased susceptibility to peroxide and increase in reactive oxygen species, indicating the importance of functional mitochondria in controlling Hog1-mediated oxidative stress response. Mitochondrial phospholipid levels were also altered as indicated by an increase in phosphatidylserine and phosphatidylethanolamine and decrease in phosphatidylcholine in fzo1Δ/Δ cells. Collectively, these findings reinforce the connection between functional mitochondria and azole tolerance, oxidant-mediated stress, and iron homeostasis in C. albicans.

INTRODUCTION

Mitochondrial biogenesis is crucial for a eukaryotic cell and involves replication of the mitochondrial genome, lipid synthesis for assembling the mitochondrial membranes, and import of nuclear encoded proteins into this organelle (1, 2). The expression of nuclear and mitochondrial encoded genes needs to be coordinated and is an essential feature of eukaryotic cells. In Saccharomyces cerevisiae, cells that have lost the mitochondrial genome (referred to as ρ⁰ cells) induce the retrograde (mitochondrial-to-nuclear signaling) and pleiotropic drug resistance (Pdr) pathways. The latter involves transcriptional induction of multidrug resistance genes coding for efflux pumps, such as PDR5, SNQ2, and YOR1, by the transcription factor (TF), Pdr3p (3). A recent study pertaining to the signal that activates the Pdr pathway in ρ⁰ cells shows that mislocalization of PSD1 (phosphatidylserine decarboxylase), the mitochondrial phosphatidylethanolamine (PE) biosynthesis enzyme, is one of the signals that is detected by Pdr3p, which in turn activates the plasma membrane localized major efflux pump, Pdr5p, leading to drug resistance in S. cerevisiae (4).

The emergence of inherently drug-resistant fungi, which are causative agents for global infectious diseases, is on the rise. Resistance to azoles, the most commonly used antifungal, occurs as a result of multiple mechanisms operating in combination in a single isolate (5). Several studies have reported the association of mitochondrial dysfunction with decreased susceptibility to fluconazole in Candida glabrata, as in the case of S. cerevisiae. The altered drug susceptibility in a set of mutants, obtained from screening a transposon mutant library in C. glabrata, is associated with genes involved in retrograde signaling or in mitochondrial biogenesis (6). A link between mitochondrial deficiency, decreased susceptibility to azoles, and increased virulence has been observed in a C. glabrata strain, BPY41 (7). The strains BPY40 (azole sensitive) and BPY41 (azole resistant) were isolated from a patient undergoing azole therapy, and the observed decreased susceptibility to azoles is attributed to the upregulation of drug efflux pumps encoded by CgCDR1, CgCDR2, and CgSNQ2 genes of the ABC transporter superfamily. Microarray profiling on BPY41 demonstrate an upregulation in genes associated with oxidoreductive metabolism and stress response consistent with dysfunctional mitochondria. That study (7) shows that defects in mitochondrial function confer selective advantage to the strain by increasing its fitness in the host.

Parallel studies on the link between drug resistance, virulence, and mitochondrial dysfunction in the opportunistic human pathogenic fungus C. albicans are limited. A petite mutant P5, recovered by serial passaging in mice spleen displays a decrease in susceptibility to fluconazole and voriconazole (not affected by itraconazole and ketoconazole) and uncoupled oxidative phosphorylation (8). A connection between dysfunctional mitochondria, drug susceptibility, cell wall integrity, phospholipid homeostasis, and virulence is addressed in C. albicans by generating mutants of (i) GOA1 (Growth and Oxidant Adaptation), a protein that translocates from the cytosol to the mitochondria during oxidant and osmotic stress (9), and (ii) SAM37 (Sorting and Assembly Machinery), a protein localized on the mitochondrial outer membrane (10). Both mutants differ in their susceptibility to various antifungals tested. The deletion of GOA1 renders the cells susceptible to azoles, oxidative stress, inhibitors of complex I of the electron transport chain and to salicylhydroxamic acid, which inhibits the alternate oxidase pathway (11, 12). Moreover, the goa1Δ/Δ strain is more susceptible to killing by neutrophils and is avirulent in a systemic model of candidiasis (9, 13). Sam37p analysis shows a link between mitochondrial function, cell wall integrity, and phospholipid homeostasis with no effect on azole susceptibility. The sam37Δ/Δ strain displays fitness defects in vitro and in vivo and is essential for virulence (10). The molecular mechanisms linking mitochondria to the aforementioned phenotypes are not well understood in C. albicans.

In the present study, we sought to further elucidate the effects of dysfunctional mitochondria in C. albicans. In S. cerevisiae, ρ⁰ cells are obtained by deleting FZO1, a gene encoding a GTPase with a transmembrane domain involved in mitochondrial fusion, resulting in the fragmentation of the tubular mitochondrial network (14, 15). Since mitochondrial biogenesis is a sum of fusion and fission processes, an imbalance in these events results in either small distinct mitochondria or in extended interconnected networks (16). Given the role of ScFZO1 in mitochondrial biogenesis, we targeted CaFZO1 in our study to generate a mutant that would be blocked at the step of biogenesis of the organelle. Moreover, FZO1 is unique to the fungal kingdom, with no known significant homolog in human or murine genomes (the functional orthologs MFN1 and MFN2 are not homologous with FZO1), adding to its potential value as a target for antifungal drug development (17). Thereafter, we set out to test the effect of deleting CaFZO1 on mitochondrial functions. We show that deletion of FZO1 causes impaired biogenesis of mitochondria, leading to aberrant mitochondrial morphology, loss of mitochondrial DNA (mtDNA), and reduced membrane potential across the mitochondrial membrane. Deletion of FZO1 affects susceptibility to azole antifungals and peroxide by virtue of reduced activity of the Cdr1p efflux pump and impaired activation of the HOG pathway, respectively. Network modeling based on the transcription profiling data obtained from the fzo1Δ/Δ cells shows coregulation of clusters of genes associated with specific biological processes. The data link mitochondria with altered iron homeostasis and shows that, as a consequence, fzo1Δ/Δ is compromised in total ergosterol levels. We also correlate mitochondrial dynamics and phospholipid homeostasis by analyzing the steady-state levels of mitochondrial phospholipids. Furthermore, deletion of FZO1 in a matched-pair of clinical isolates renders the azole-resistant (AR) isolate moderately susceptible to azoles, indicating the potential of targeting mitochondria to reverse drug resistance. Taken together, our results highlight the pleiotropic effects that occur in a cell with dysfunctional mitochondria and validate FZO1 as a potential drug target.

MATERIALS AND METHODS

Strains, chemicals, and growth conditions.

The C. albicans strains used in the present study are listed in Table 1. The strains were maintained as frozen stocks and propagated at 30°C on the following media. YNB-glucose medium consists of yeast nitrogen base (YNB) without amino acids (Difco) and glucose (0.67% YNB, 2% glucose, 2.5% agar). YEPD (1% yeast extract, 2% peptone, 2% glucose, 2.5% agar) liquid medium and agar plates containing 200 μg of nourseothricin (Werner Bioreagants) ml⁻¹ were used to select for deletion mutants. To obtain nourseothricin-sensitive derivatives of transformants, strains were grown in YPM (1% yeast extract, 2% peptone, 2% maltose) for 8 h and plated on 25 μg ml⁻¹ nourseothricin. RPMI 1640 (containing l-glutamine, without bicarbonate) was prepared at 10.4 g liter⁻¹, buffered with 0.165 mol of MOPS (3-[N-morpholino] propanesulfonic acid) liter⁻¹, and the pH was adjusted to 7.0 using 1 mol of sodium hydroxide liter⁻¹. The following supplements, fluconazole (Sigma), ketoconazole (Sigma), itraconazole (Sigma), amphotericin B (Sigma), MitoTracker Green FM (MGFM; Molecular Probes/Invitrogen), 4′,6-diamidino-2-phenylindole (DAPI; Molecular Probes/Invitrogen), 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl benzimidazolylcarbocyanine iodide or JC-1 (Molecular Probes/Invitrogen), 2,7-dichlorofluoresceindiacetate (Sigma) and rhodamine 6G (R6G; Sigma) were added to the medium or buffer at the concentrations described. Ergosterol standard and dthe erivatizing agent N,O-Bis (trimethylsilyl) trifluoroacetamide with trimethylchlorosilane (TCMS) were purchased from Sigma.

Table 1.

C. albicans strains used in this study

Strain	Parent	Genotype	Source or reference
SC5314		Wild type	89
ET10	SC5314	FZO1/fzo1Δ::SAT1-FLIP	This study
ET11	ET10	FZO1/fzo1Δ::FRT	This study
ET12	ET11	fzo1Δ::SAT1-FLIP/fzo1Δ::FRT	This study
ET13	ET12	fzo1Δ::FRT/fzo1Δ::FRT	This study
ET14	ET13	fzo1Δ::FRT/fzo1Δ::FZO1-SAT	This study
GU4	Clinical isolate (fluconazole susceptible)	Wild type	71
ET16	GU4	FZO1/fzo1Δ::SAT1-FLIP	This study
ET17	ET16	FZO1/fzo1Δ::FRT	This study
ET18	ET17	fzo1Δ::SAT1-FLIP/fzo1Δ::FRT	This study
ET19	ET18	fzo1Δ::FRT/fzo1Δ::FRT	This study
ET20	ET19	fzo1Δ::FRT/fzo1Δ::FZO1-SAT1-FLIP	This study
GU5	Clinical isolate (fluconazole resistant)	Wild type	71
ET21	GU5	FZO1/fzo1Δ::SAT1-FLIP	This study
ET22	ET21	FZO1/fzo1Δ::FRT	This study
ET23	ET22	fzo1Δ::SAT1-FLIP/fzo1Δ::FRT	This study
ET24	ET23	fzo1Δ::FRT/fzo1Δ::FRT	This study
ET25	ET24	fzo1Δ::FRT/fzo1Δ::FZO1-SAT1-FLIP	This study
ET26	SC5314	PCDR1-CDR1-GFP integrated at the CDR1 locus	This study
ET27	ET13	PCDR1-CDR1-GFP integrated at the CDR1 locus	This study

Strain construction.

The first and second allele of the FZO1 gene was disrupted by using the SAT1 flipper in the plasmid pSFS2 (18). For the FZO1 disruption construct, a 1,000-bp 5′ upstream noncoding region (NCR) of FZO1 (5′FZO1^NCR) was amplified from SC5314 genomic DNA with the primers FP2 and FP3 (see Table S1 in the supplemental material), which introduced KpnI and XhoI restriction sites. A 500-bp 3′ FZO1^NCR was amplified with primers FP4 and FP5, which introduced NotI and SacI sites. All amplicons (5′ FZO1^NCR and 3′ FZO1^NCR) were docked in pGEM-T Easy, from which they were digested and cloned into the 5′ and 3′ ends of the CaSAT1-FLIP cassette, respectively, using the mentioned enzymes. This procedure created the FZO1 knockout construct plasmid pET1 (Table 2 and Fig. 1A), which was cut with KpnI and SacI to release the 5.7-kb disruption construct. The wild-type SC5314 strain was electroporated with the disruption construct. The CaSAT1 construct was flipped out from FZO1/fzo1Δ::SAT1-FLIP strain before disruption of the second allele. For the disruption of the second allele, 3′ end of pET1 was replaced by 500 bp of the FZO1 open reading frame (ORF; amplified by primers FP6 and FP7) with NotI and SacI restriction sites, generating pET2 (Table 2). The FZO1 reconstitution construct was made by amplifying a 2.9-kb fragment containing the FZO1 ORF and the 5′ NCR by using the primers FP8 and FP9 that introduced KpnI and HindIII sites. This fragment was ligated into the KpnI-HindIII-digested pET1, which already contained the 3′ FZO1^NCR. This procedure resulted in the FZO1 reconstitution plasmid pET3 (Fig. 1B).

Table 2.

Plasmids used in this study

Plasmid	Description	Source or reference
pSFS2B	SAT1 flipper carrying nourseothricin resistance gene	18
pET1	pSFS2B flanked 5′ and 3′ FZO1NCR for disruption of first allele of FZO1	This study
pET2	pSFS2B flanked 5′ FZO1NCR and 3′ FZO1ORF for disruption of second allele of FZO1	This study
pET3	FZO1 reconstitution construct	This study
pCPG2	Plasmid harboring CDR1-GFP fusion for CDR1 locus integration	45

Fig 1.

(A) Schematic representation of disruption strategy. (B) FZO1 reconstitution construct used to integrate FZO1 in the fzo1Δ/Δ mutant. (C) Southern hybridization showing genomic configuration of FZO1 in the wild-type (WT) locus and its deletion derivatives. Genomic DNA from strains was digested with PstI. Lane 1, wild type (SC5314); lane 2, FZO1/fzo1Δ::SAT1-FLIP; lane 2, FZO1/fzo1Δ::FRT; lane 3, fzo1Δ::SAT1-FLIP/fzo1Δ::FRT; lane 4, fzo1Δ::FRT/fzo1Δ::FRT; lane 5, fzo1Δ::FRT/fzo1Δ::FZO1-SAT1-FLIP. (D) Growth curve of wild type, null mutant, and the reconstituted strain in YNB-glucose. Overnight cultures were diluted to an OD₆₀₀ of 0.1, and growth was monitored by measuring the OD of the cultures over time. The doubling time of each strain in YNB-glucose is mentioned. All values are means ± the standard deviations (SD) of three independent experiments performed in duplicates. (E) The strains were incubated at 30°C, grown to early log phase, and stained with 20 nM MGFM for 30 min. The cells were viewed by confocal microscopy to assess the mitochondrial morphology.

Southern blot analysis.

The cells were grown in 10 ml of YEPD at 30°C overnight. The culture was harvested and genomic DNA was extracted. Then, 10 μg of genomic DNA was digested with PstI, separated on a 1% agarose gel, transferred onto a nylon membrane, and fixed by UV cross-linking. The gel-purified 500-bp 3′ FZO1^NCR fragment was used as a probe. The probe was labeled with [³²P]dATP. All blots were hybridized at 65°C in a solution containing 0.5 M NaH₂PO₄ (pH 7.2), 7% sodium dodecyl sulfate (SDS), and 1 mM EDTA. After hybridization, the membranes were washed thrice with 2× SSC (1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate) and 0.1% SDS. All membranes were imaged by using a Fujifilm phosphorimager (Amersham Biosciences).

Microscopy.

For confocal microscopy, an Olympus FluoView FV1000 microscope (×100 oil immersion objective lens) was used, and photographs were processed with the Olympus FV110A SW 1.7 viewer software. For visualizing mitochondrial morphology, nuclear and mitochondrial DNA, and green fluorescent protein (GFP)-tagged protein, cells were grown to mid-log phase and stained with MGFM and DAPI. MGFM and DAPI were added to concentrations of 20 nM and 1 μg ml⁻¹, respectively, and incubated for 20 min for staining. After staining, the cells were washed thrice and resuspended in phosphate-buffered saline (PBS). The fluorescence excitation/emission values were 490/516 nm and 358/461 nm for MGFM and DAPI, respectively.

Antifungal susceptibility tests.

Strains grown overnight in YNB-glucose were diluted in 0.9% saline solution. Then, 5-μl portions of four dilutions (5 × 10³to 5 × 10⁵ cells) were spotted onto YNB-glucose containing indicated drugs. Plates were photographed after 48 h at 30°C. The MIC was determined by broth microdilution methods described in Clinical and Laboratory Standards Institute guidelines. The diluted cell suspensions (10⁴ cells ml⁻¹) were added to the wells of round-bottom 96-well microtiter plates containing equal volumes of RPMI 1640 (buffered with 0.165 mol of MOPS liter⁻¹) and serially diluted concentrations of drugs. The plates were incubated at 37°C for 48 h. The growth was evaluated by reading the optical density at 600 nm (OD₆₀₀) in a microplate reader and MIC₈₀ is defined as the lowest drug concentration that gave 80% inhibition of growth compared to the growth of the drug-free controls.

Efflux of R6G.

Approximately 10⁷ yeast cells from each overnight culture were inoculated into 50 ml of YEPD medium and allowed to grow for 5 h. The cells were pelleted, washed twice with PBS buffer (pH 7.0), and resuspended in PBS to 2% cell suspension. R6G was added at a final concentration of 10 μM. Cell suspensions were incubated at 30°C with shaking (200 rpm) for 3 h under glucose starvation conditions. The de-energized cells were then washed and resuspended again in PBS at a 2% cell suspension. At 10-min intervals, a portion (1 ml) of the cells was removed and centrifuged, and the absorption of the supernatants was measured at 527 nm. Energy-dependent efflux was measured after the addition of 2% glucose. Glucose-free controls were included in all experiments. Effluxed R6G was calculated from a standard concentration curve of R6G.

Flow cytometry assays.

Overnight cultures grown at 30°C were introduced into fresh YNB-glucose/glycerol/galactose and incubated for 6 to 8 h. The cells were washed twice with PBS and treated with 5 μM JC-1 or 10 μM DCFDA (2′,7′-dichlorofluorescein diacetate). Samples were maintained for 30 min at 30°C in the dark and washed thrice with PBS, and the fluorescence was measured with a Becton Dickinson flow cytometer. An unstained sample was used as a control. For JC-1 experiment, control cells were incubated with sodium azide (20 mM) and oligomycin (0.5 μg ml⁻¹) for 30 min prior to staining to induce a decrease in the membrane potential. Control cells were treated with 10 mM H₂O₂ for 30 min to increase reactive oxygen species (ROS) production prior to DCFDA staining. Flow cytometry was performed using a FACSCalibur flow cytometer (Becton Dickinson Immunocytometry Systems, San Jose, CA) equipped with an argon laser emitting at 488 nm. Fluorescence was measured on the FL1 fluorescence channel equipped with a 530-nm band-pass filter for GFP and DCFDA staining. The fluorescence intensities at FL-1 and FL-2 (red fluorescence, 595 nm) were recorded for JC-1 staining. A total of 10,000 events were counted. The data were analyzed using CellQuest V software.

Ergosterol estimation.

Briefly, a single C. albicans colony from an overnight YNB-glucose plate culture was used to inoculate 100 ml of YNB-glucose broth. The cultures were incubated for 16 h with shaking at 30°C. The cells were harvested by centrifugation at 6,000 rpm for 5 min and washed twice with sterile-distilled water. The net wet weight of the cell pellet was determined. Equal cell weights were resuspended in 10 ml of methanol, and cells were broken with glass beads by ultrasonication with a Branson Digital Sonifier. Two volumes of chloroform were added, and the suspension was stirred for 2 h and filtered. To this extract, 0.2 volume of 0.9% saline was added. After the aqueous layer was aspirated off, the lipids were dried down under nitrogen, dissolved in 100 μl of TCMS, and incubated at 82°C for 60 min. After cooling, it was dried under nitrogen and net lipid weight was measured. Derivatized sterols were then resuspended in chloroform and used for gas chromatography-mass spectrometry (GC-MS) quantification. Ergosterol standards of known concentrations, also derivatized with TCMS, and the sample-derivatized sterols were then run through a Restek RTX-5MS Crossbond 5% diphenyl–95% dimethyl polysilane column in a gas chromatograph-mass spectrometer (model QP2010 Plus; Shimadzu, Japan). Samples were analyzed in reference to retention times and integrated peak areas by using GCMS-QP2010 Series software. The results are presented as mg of ergosterol g of lipid weight⁻¹ according to the calibration from the standard curve of ergosterol.

Microarray analysis.

(i) For transcriptional profiling, wild-type and mutant cells were grown overnight in 10 ml of YNB-glucose at 30°C and 200 rpm, subcultured to an OD₆₀₀ of 0.1 in 100 ml of YNB-glucose medium, and grown for 7 h at 30°C and 200 rpm. RNA was extracted from three biological replicates of both strains using an RNeasy minikit (Qiagen). The RNA samples of three biological replicates of each strain were used to perform the microarray experiments. The labeled cRNA samples were hybridized on to a custom C. albicans 8×15K microarray designed by Genotypic Technology Pvt., Ltd., Bangalore, India (Amadid no. 026377). Cy3-labeled samples (600 ng) were fragmented and hybridized using an Agilent gene expression hybridization kit (catalog no. 5188-5242). Hybridization was carried out in Agilent SureHyb chambers at 65°C for 16 h, and hybridized slides were scanned using a microarray scanner (G2505C; Agilent) at 5-μm resolution. Data extraction from images was done using Agilent's Feature Extraction software (v10.5.1.1). (ii) For data analysis, all microarray data were analyzed using the GeneSpring GX v12.0 from Agilent Technologies (Santa Clara, CA). Preprocessing of the data was carried out by calculating the 50th percentile for the intrasample value and normalizing that to the median of all samples for intersample normalization. Replicate sample reproducibility was performed using principle component analysis to understand the biological variation within replicate samples. (iii) For differential expression analysis, genes that were differentially expressed in mutant samples in compared to wild-type samples were evaluated by using the Volcano plot method. Genes that were ≥2-fold up- or downregulated in mutant cells with a P value of <0.05 were considered differentially expressed. An unpaired Student t test was used to calculate the P value. Differentially expressed genes, along with the samples, were clustered using unsupervised hierarchical clustering method with the Pearson uncentered algorithm and the average linkage rule. (iv) Gene ontology (GO) analysis was carried out using the GO Term Finder at the CGD (Candida Genome Database; http://www.candidagenome.org/cgi-bin/GO/goTermFinder). Upregulated and downregulated genes were analyzed separately. (v) For biological analysis network modeling, one or more differentially expressed genes that were associated with significantly enriched processes were clustered together to create a network. Over-representation analysis by Cytoscape v8.0 was performed to identify genes that are key regulatory nodes activated or repressed in the mutant.

Quantitative real-time PCR.

C. albicans strains were grown overnight in YNB-glucose, subcultured from a starting OD₆₀₀ of 0.1 in fresh YNB-glucose (with or without 100 μM FeCl₃), and incubated at 30°C for 7 h. Total RNA, isolated using the RNeasy minikit (Qiagen), was treated with DNase I (Fermentas Life Sciences) to remove contaminating DNA. cDNA was synthesized with a RevertAid H Minus First Strand cDNA synthesis kit (Fermentas Life Sciences) according to the manufacturer's protocol. Real-time PCRs were performed in a volume of 25 μl using the Thermo Scientific Maxima SYBR green mix in a 96-well plate. For the relative quantification of gene expression, the comparative threshold cycle (C_T) method was used, where the fold change was determined as 2^−ΔΔCT (19). ACT1 was used as the internal control, and the transcript level of the gene of interest was normalized to the ACT1 levels. The quantitative real-time PCR (qPCR) primers used in the present study were designed by Primer Express 3.0 and are listed in Table S1 in the supplemental material.

Steady-state phospholipid analysis.

Phospholipids were extracted from crude mitochondrial fractions and separated by thin-layer chromatography (TLC) as described previously (20, 21). Briefly, starter cultures were diluted to an OD₆₀₀ of 0.2 in YNB-glucose supplemented with 10 μCi of ³²P_i ml⁻¹ and grown at 30°C for 24 h. After a wash with H₂O, the yeast pellets were resuspended in breaking buffer and disintegrated by vortexing with glass beads for 30 min at 4°C. Phospholipids from equal amounts of labeled crude mitochondria, as determined by liquid scintillation, were extracted with a 2:1 chloroform-methanol by vortexing at room temperature for 1 h. After phase separation, the lower organic phase was transferred to a new borosilicate tube and dried down under a stream of liquid nitrogen. Chloroform-resuspended samples were loaded onto silica gel TLC plates and resolved. Statistical comparisons were performed using SigmaPlot 11 software (Systat Software, Inc.).

Protein extracts and immunoblot analysis.

For kinetics assays, overnight cultures were diluted in fresh YEPD medium to an A₆₀₀ of 0.05 and grown until they reached an A₆₀₀ of 1 at 37°C and 200 rpm. Samples were treated with 10 mM hydrogen peroxide before they were recovered. The procedures used for cell collection, lysis, protein extraction, fractionation by SDS-PAGE, and transfer to nitrocellulose membranes have been previously described (22). Anti-phospho-p44/p42 mitogen-activated protein kinase (MAPK; Thr202/Tyr204) antibody (New England BioLabs) was used to detect dually phosphorylated Mkc1 and Cek1 MAPKs; phospho-p38 MAP kinase (Thr180/Tyr182) 28B10 monoclonal antibody (Cell Signaling Technology, Inc.) and ScHog1 polyclonal antibody (Santa Cruz Biotechnology) were used to detect the phosphorylated Hog1 and Hog1 proteins, respectively. Western blots were developed according to the manufacturer's conditions using the Hybond ECL kit (Amersham Pharmacia Biotech). To equalize the amounts of protein loaded, samples were analyzed by measuring the A₂₈₀ and then by Coomassie staining. In addition, the anti-ScHog1 signal was used both as an internal control in experiments concerning activation of CaHog1 and as an additional loading control when other MAPKs were analyzed; this antibody was sometimes mixed with compatible antibodies as internal controls.

Accession number.

The microarray data can be accessed under GEO accession number GSE46003.

RESULTS

Growth and mitochondrial parameters are affected in the absence of FZO1.

Deletion of FZO1 was accomplished by the SAT-flipper strategy (18) in the wild-type strain SC5314 (Fig. 1A). FZO1 was reconstituted into its native locus in the fzo1Δ/Δ strain (Fig. 1B). Each step of gene deletion and reconstitution was confirmed by Southern blotting (Fig. 1C). To determine the effect of loss of FZO1 on growth characteristics, the wild-type, fzo1Δ/Δ, and reconstituted strains were spotted on rich YEPD and synthetic YNB-glucose media. The mutant displayed a fitness defect on both media, the defect being more pronounced on the YNB-glucose medium (see Fig. S1 in the supplemental material [growth controls]). In liquid synthetic media, while the wild type had a mean doubling time of 1.56 ± 0.1, the fzo1Δ/Δ mutant had a significant higher mean doubling time of 1.97 ± 0.1 (Fig. 1D). The reconstituted strain had a mean doubling time of 1.57 ± 0.05, similar to that of the wild type. This defect was observed in two independent clones of the null mutant.

We hypothesized that the slow-growth phenotype of the mutant could be associated with impaired mitochondrial parameters such as morphology, genome maintenance and function. The length, shape, size, and number of mitochondria are controlled by fusion and fission events. At steady state, the frequencies of fusion and fission events are balanced to maintain the overall morphology of the mitochondrial population. When this balance is experimentally perturbed, transitions in mitochondrial shape can occur. Although the loss of FZO1 renders S. cerevisiae incompetent in the fusion of mitochondria, its role in C. albicans has not been studied. Since mitochondrial fragmentation is indicative of reduced or blocked fusion activity, we tested the C. albicans fzo1Δ/Δ mutant for the presence of fragmented mitochondria by using the dye MGFM. In the wild type, mitochondria were visualized as tubular networks spanning the periphery of the cell, whereas the mutant displayed clumps or aggregated masses in the cytoplasm, likely due to continuing fission events (Fig. 1E). The reconstituted strain showed mitochondrial morphology similar to that of the wild type, indicating that the aberration is reversible (Fig. 1E).

An additional secondary phenotype that is observed in cells with fragmented mitochondria is their tendency to lose the mitochondrial genome (mtDNA), leading to respiratory incompetence (14, 15, 23, 24). Therefore, we tested the mutant (i) for loss of mtDNA by DAPI staining and (ii) for growth on nonfermentable carbon sources, such as glycerol and ethanol. The data for DAPI staining revealed that 94% of the mutant cells grown in either YEPD or YNB-glucose were devoid of mtDNA, compared to the wild type (Fig. 2A and B). The reconstituted strain showed mtDNA similar to the wild type, indicating that FZO1 is required for mtDNA stability. Loss of mtDNA was also reflected in the inability of the mutant to grow in the presence of nonfermentable carbon sources. Interestingly, the mutant was able to grow on YP supplemented with glycerol and ethanol, albeit at a slow rate, but was unable to grow on YNB containing glycerol and ethanol (both at 2%) (Fig. 2A and see Fig. S1 in the supplemental material). In addition, we observed a significant reduction in the transcript levels of the mtDNA-encoded genes ATP6, COX2, COX3, NAD1, and NAD2 (Fig. 2C). This observation indicates that while the majority of fzo1Δ/Δ cells are devoid of mtDNA, a few cells that retain wild-type mtDNA not only restore the mitochondrial morphology to wild type after reconstitution but also account for the presence of transcripts of mtDNA-encoded genes in the mutant.

Fig 2.

(A) The strains were incubated at 30°C, grown to early log phase, and stained with 1 μg of DAPI ml⁻¹ for 20 min. Cells were viewed by using confocal microscopy. The wild-type strain showed a larger nuclear DNA (denoted by “N”), along with several smaller mtDNA nucleoids (denoted by “M”). The growth of the strains on nonfermentable carbon sources (glycerol + ethanol) is indicated as follows: +++, optimum growth; +; slow growth; –, no growth. (B) Quantitation of cells containing mtDNA. Approximately 500 cells were visualized in each experiment. Values are means ± the SD (n = 3). (C) mtDNA loss was confirmed by qPCR of the mtDNA genes ATP6, COX2, COX3, NAD1, and NAD2. The relative transcript level was calculated by using 2^−ΔΔCT, normalized to ACT1 (endogenous control). The dashed line at 1.0 indicates wild type used as a calibrator for the calculations. (D) Altered mitochondrial membrane potential demonstrated as the ratio of mean fluorescence (FL-2/FL-1) in the wild type and null mutant grown to the log phase in YNB-glycerol (2%) after staining with JC-1. Wild-type cells were also treated with sodium azide as a control. The inset shows the FL-2/FL-1 ratio in the wild type and null mutant grown in YNB-glucose. *, P < 0.01 (Student t test).

In order to assess the effect of deletion of FZO1 on mitochondrial functions, we analyzed the membrane potential across the mitochondrial membrane using the cationic dye JC-1. JC-1 discriminates energized and de-energized mitochondria in a potential dependent manner, where red fluorescence increases in response to high mitochondrial membrane potential. Hence, as a measure of mitochondrial activity, the accumulation of the dye has been used in C. albicans to quantify the mitochondrial membrane potential (25, 26). Cells grown in YNB-glucose or YNB-glycerol were stained with JC-1 prior to fluorescence detection by flow cytometry and mitochondrial polarization was measured as an increase in the red (J-aggregated)/green (monomer) fluorescence intensity ratio (FL-2/FL-1). Wild-type cells grown in YNB-glucose displayed green fluorescence, which is suggestive of fermentative metabolism. However, the wild-type cells upon growth in YNB-glycerol showed an increase in red/green ratio, indicating polarization (aggregation of the dye) of the mitochondrial membrane due to actively respiring mitochondria (Fig. 2D). There was no difference in the red/green ratio between the wild type and the mutant grown in YNB-glucose (Fig. 2D, inset). The deletion of FZO1 lowered the red/green ratio relative to the wild type, similar to that observed in cells incubated with mitochondrial inhibitor cocktail (oligomycin and sodium azide) (Fig. 2D). In lieu of the above data, we surmise that the fzo1Δ/Δ mutant (i) harbors aberrant mitochondria, (ii) has perturbed mitochondrial membrane potential, and (iii) is unable to utilize nonfermentable carbon sources. Further, the latter two phenotypes likely stem from the absence of mtDNA in the fzo1Δ/Δ mutant, which encodes several subunits of the oxidative phosphorylation complexes.

Transcriptome analysis.

In order to determine the global consequences of dysfunctional mitochondria, we compared the mRNA profiles of wild-type and fzo1Δ/Δ cells in synthetic defined media. A total of 649 (78%) genes were downregulated, and 179 (22%) genes were upregulated ≥2-fold, with a P value of ≤0.05. Significant gene clusters were identified that displayed differential expression between the wild type and the mutant (Table 3). Genes related to mitochondrial function (23%) topped the list of genes that were significantly upregulated in the mutant, followed by genes that are associated with transport (11%), ribosome biogenesis (11%), and response to drug (7%) (Fig. 3A). In addition, 13 genes associated with iron assimilation and/or transport were upregulated in the mutant (5%), along with genes involved in response to stress (6%) (Fig. 3A and C). The highest upregulated gene in the transcriptome profiling was AOX2 (Alternative Oxidase 2) that is involved in the cyanide-resistant respiratory pathway in C. albicans (27), followed by the multidrug transporter, MDR1, known to be involved in fluconazole tolerance (Fig. 4B) (28).

Table 3.

Functional categories of C. albicans genes whose transcript levels in a fzo1Δ/Δ strain are >2-fold up- and downregulated (P ≤ 0.05)

Category and systematic name	Genea	Function	Fold change
Mitochondrial functions
orf19.4773	AOX2*	Alternate oxidase involved in cyanide resistant respiratory pathway	73
orf19.3700	TOM70	Role in protein import into mitochondrial inner membrane and matrix	2.4
orf19.3691	TIM21	Translocase of the inner mitochondrial membrane (TIM23 complex) involved in protein import into mitochondria	2.4
orf19.5419	ATP5	Subunit of F0-F1 ATP synthase	−2.8
CaalfMp07	ATP8	Subunit of F0-F1 ATP synthase	−5.6
orf19.5491.1	ATP14	Subunit of F0-F1 ATP synthase	−3.0
orf19.7678	ATP16	Subunit of F0-F1 ATP synthase	−4.3
orf19.7509.1	ATP17	Subunit of F0-F1 ATP synthase	−5.6
orf19.2066.1	ATP18	Subunit of F0-F1 ATP synthase	−4.2
orf19.5231.2	ATP19	Subunit of F0-F1 ATP synthase	−3.5
orf19.3757	ATP20	Subunit of F0-F1 ATP synthase	−2.8
orf19.5653	ATP2	Subunit of F0-F1 ATP synthase	−2.5
CaalfMp08	COX1	Subunit of cytochrome c oxidase	−7.8
orf19.1471	COX4	Putative cytochrome c oxidase subunit IV	−2.5
orf19.4759	COX5	Cytochrome oxidase subunit V	−2.6
orf19.873.1	COX6	Putative cytochrome c oxidase	−2.5
orf19.5213.1	COX8	Putative cytochrome c oxidase	−5
CaalfMp09	NAD2*	Subunit of NADH:ubiquinone dehydrogenase	−15
CaalfMp10	NAD3	Subunit of NADH:ubiquinone dehydrogenase	−7.2
CaalfMp12	NAD4L	Subunit of NADH:ubiquinone dehydrogenase	−6.7
Carbohydrate metabolism
orf19.3982	MAL32	Maltose alpha-glucosidase activity	2.9
orf 19.3483		Involved in glycerol metabolic process	2.5
orf19.4618	FBA1	Putative fructose-bisphosphate aldolase	−3.0
orf19.691	GPD2	Glycerol 3-P dehydrogenase	−2.3
orf19.903	GPM1	Phosphoglycerate mutase	−2.9
orf19.542	HXK2	Hexokinase II	−2.6
orf19.4602	MDH1-1	Predicted malate dehydrogenase precursor	−2.5
orf19.3097	PDA1	Putative pyruvate dehydrogenase alpha chain	−2.9
orf19.2877	PDC11	Putative pyruvate decarboxylase	−3.4
orf19.6540	PFK2	Phosphofructokinase	−2.5
orf19.3651	PGK1	Phosphoglycerate kinase	−2.9
orf19.704	SOL3	Putative 6-phosphogluconolactonase	−2.7
orf19.6814	TDH3	NAD-linked glyceraldehyde-3-phosphate dehydrogenase	−2.6
orf19.6745	TPI1	Triose-phosphate isomerase	−2.9
orf19.1480		Protein described as succinate dehydrogenase, enzyme of citric acid cycle	−2.3
Transport/transporter activity
orf19.5604	MDR1*	Multidrug efflux pump from major facilitator transporter superfamily	23
orf19.5759	SNQ2	Protein similar to S. cerevisiae Snq2p transporter; member of PDR subfamily of ABC family	2.5
orf19.1783	YOR1	Protein similar to S. cerevisiae Yor1p, which is a plasma membrane transporter of the ATP-binding cassette (ABC) family	2.7
orf19.4527	HGT1	Putative glucose transporter of the major facilitator superfamily	4.1
orf19.7148	TPO2	Putative polyamine transport protein	2.8
orf19.5672	MEP2	Ammonium permease	−2.5
orf19.1585	ZRT2	Predicted zinc transporter	−4.4
orf19.6296	VPS22/SNF8	ESCRT-II complex protein with a role in multivesicular body trafficking	−2.2
Lipid metabolism
orf19.4122	TES2	Acyl-CoA hydrolase activity, role in fatty acid beta-oxidation	2.5
orf19.3483		Putative phosphatidylglycerol phospholipase C	2.5
orf19.406	ERG1*	Squaleneepoxidase	−3.1
orf19.767	ERG3*	C-5 sterol desaturase	−4.0
orf19.922	ERG11*	Lanosterol 14-α-demethylase	−3.1
orf19.5379	ERG4	Protein similar to sterol C-24 reductase	−2.3
orf19.3732	ERG25	Putative C-4 methyl sterol oxidase	−3.1
orf19.979	FAS1	Beta subunit of fatty acid synthase	−2.0
orf19.5949	FAS2	Alpha subunit fatty acid synthase	−2.1
orf19.3822	SCS7	Putative ceramide hydroxylase	−2.7
orf19.5818	SUR2	Putative ceramide hydroxylase of sphingolipid biosynthesis	−2.4
orf19.6105	MVD	Mevalonate diphosphate decarboxylase (homolog of ScERG19)	−2.1
orf19.391	UPC2	Zn2-Cys6 transcriptional regulator of ergosterol biosynthetic genes and sterol uptake	−4.6
Cell wall organization
orf19.5741	ALS1	Adhesin, cell surface glycoproteins	5.2
orf19.7114	CSA1	Cell surface antigen present on elongating hyphae and buds	2.4
orf19.3966	CRH12	Putative cell wall protein	−2.6
orf19.7436.1	ECM15	Predicted role in cell wall organization	−3.1
orf19.4887	ECM21	Protein similar to S. cerevisiae Ecm21p (possible role in cell wall)	−2.6
orf19.3010.1	ECM33	GPI-anchored cell wall protein	−2.7
orf19.4035	PGA4	GPI-anchored cell surface protein	−3.1
orf19.968	PGA14	GPI-anchored cell surface protein	−2.0
orf19.2475	PGA26	GPI-anchored cell surface protein	−2.4
orf19.5302	PGA31	GPI-anchored cell surface protein	−2.4
orf19.1911	PGA52	GPI-anchored cell surface protein	−2.4
orf19.2765	PGA62	GPI-anchored cell surface protein	−3.0
orf19.4109	PMT4	Protein mannosyltransferase required for normal cell wall composition and virulence	−2.0
orf19.1390	PMI1	Phosphomannose isomerase; cell wall biosynthesis enzyme	−2.1
orf19.6081	PHR2	Glycosidase; role in cell wall structure	−2.3
orf19.7218	RBE1	Cell wall protein	−4.1
Iron assimilation/transport
orf19.4328	CCC2*	Copper-transporting P-type ATPase required for iron assimilation	3.1
orf19.2179	SIT1*	Transporter of ferrichrome siderophores	7.9
orf19.5636	RBT5*	GPI-anchored cell wall protein involved in hemoglobin utilization	6.6
orf19.1932	FRE5*	Ferric reductase	22
orf19.1264	FRE2*	Putative oxidoreductase	1.29
orf19.1930	FRE31*	Ferric reductase	6.52
orf19.4647	HAP32*	CCAAT-binding transcription factor that regulates respiration	12.4
orf19.1267.1		Role in iron-sulfur cluster assembly	2.3
orf19.2803	HEM13	Coproporphyrinogen III oxidase	−3.6
orf19.4747	HEM14	Putative protoporphyrinogen oxidase involved in heme biosynthesis	−3.3
Biotin synthesis
orf19.2591	BIO3*	Role in biotin biosynthetic process	−6.0
orf19.2590	BIO4*	Dethiobiotin synthetase	−6.4

^a*, Genes that were validated by qPCR.

Fig 3.

Functional categories are represented as the percentages of total genes that were upregulated (A) and downregulated (B). (C) Visualization of the coexpression network in fzo1Δ/Δ mutant representing the three structured distinct modules of differentially expressed genes involved in specific biological processes. Clustering was based on over-representation analysis using Cytoscape v8.0.

Fig 4.

(A) Fivefold serial dilutions of cell suspensions were spotted onto YNB-glucose plates supplemented with drugs indicated and incubated at 30°C for 48 h. (B) qPCR of CDR1, CDR2, and MDR1 transcript levels in the mutant. The fold change was calculated by using 2^−ΔΔCT, normalized to ACT1 (endogenous control), with the wild type as the calibrator. (C) Efflux of fluorescent R6G, a substrate of CDR pumps. All strains were grown overnight in YEPD, starved for 2 h in PBS, incubated with R6G (10 μM), and then transferred to PBS (pH 7). At 10 min, glucose was added to cultures, and the efflux of fluorescent rhodamine was measured subsequently for a total of 40 min. (D) Confocal micrographs of wild-type and fzo1Δ/Δ cells containing the chromosomally integrated P_CDR1-CDR1-GFP reporter fusion. The values below indicate mean fluorescence measurements ± the SD (n = 3) determined by flow cytometry. WT, wild type.

GO enrichment analysis showed significant downregulation of transport and mitochondrial genes (Table 3). The five highest categories of genes that were downregulated were associated with transport (12%), mitochondria (12%), stress response (6%), and carbohydrate metabolism (4%). Genes associated with key processes like, cell wall biosynthesis (5%), cell cycle (3%), and lipid metabolism (3%) were also listed as significantly downregulated genes (Fig. 3B).

The integrated transcriptome results indicated the existence of three specific clusters of genes that are coregulated in response to dysfunctional mitochondria in the mutant (Fig. 3C). The first and second subsets of genes contained genes involved in cell cycle and cell wall organization. The third subset of deregulated genes contains those that are involved in iron homeostasis. A minor cluster of genes involved in cell adhesion or virulence (ALS1) is also downregulated in the mutant. The resulting network also highlighted key molecules involved in cross talk of the three major biological processes represented in the network and its regulatory control. Genes such as YOX1, CBF1, ACT1, PGA26, PHR2, and PGA62 were found to be key nodes that connect deregulated biological processes such as cell adhesion, cell cycle, cell wall organization, iron assimilation/transport, transcription, and virulence (Fig. 3C).

In S. cerevisiae, the early cell cycle box (ECB) elements, which facilitate the M/G₁ specific transcription, are necessary for the expression of genes required for proceeding to the S phase and comprise of a Mcm1 binding site (29, 30). ScYOX1 is a repressor that binds to Mcm1 and prevents the ECB-mediated transcription to the M/G₁ interval of the cell cycle in the budding yeast. In C. albicans, the expression of YOX1 peaks at the G₁/S phase of the cell cycle and, unlike ScYOX1, it does not have a role in M/G₁ transition (31, 32). Deletion of CBF1 causes a severe growth defect in C. albicans (33) and has been shown to bind to the upstream region of ribosomal protein coding genes, sulfur starvation regulons, and one-quarter of the respiratory-chain-coding genes (34, 35). These data suggest transcriptional remodeling in the mutant to facilitate the reduction of energy-consuming processes such as cell division and cell wall maintenance. The expression of genes such as PGA62, PGA26, and PHR2 is modulated by iron concentrations (36), which is in line with their presence at nodes in the coregulatory network linking iron homeostasis genes with cell wall organization genes.

Noteworthy among the gene clusters was the set that contained genes involved in iron homeostasis (Fig. 3C), suggesting that multiple iron metabolism-dependent pathways were remodeled in the mutant. Genes that belong to the iron-regulon were significantly upregulated in the mutant (Table 3). Repression of genes encoding proteins of the mitochondrial respiratory chain was also observed. This set included genes for several subunits of NADH:ubiquinone dehydrogenase, F₁F_o-ATP synthase and the subunits of cytochrome c oxidase (Table 3). The genes encoding for the respiratory chain proteins are regulated by iron availability and are repressed upon iron limitation in S. cerevisiae (37, 38). HGT1 (high-affinity glucose transporter) a gene involved in glucose transport/acquisition was induced, suggesting a compensatory response. Genes of other biosynthetic pathways that are modulated by iron-availability, such as ergosterol biosynthesis, biotin synthesis (BIO3 and BIO4), ammonium uptake (MEP2), and heme metabolism were also repressed in the mutant. Thus, we surmise that the major effects of dysfunctional mitochondria was (i) a much larger set of 649 genes was downregulated and (ii) a coregulated response of genes involved in specific biological processes was triggered.

Increased azole susceptibility of fzo1Δ/Δ cells is due to decreased activity of the CDR1 efflux pump.

Mutants having aberrant mitochondrial function are known to have a decreased susceptibility to azole antifungals and polyenes in S. cerevisiae and C. glabrata (39, 40). In C. albicans, deletion of mitochondria localized GOA1 renders cells highly susceptible to azoles and cell wall damaging agents (12, 13). Considering that deletion of FZO1 in S. cerevisiae does not alter tolerance to fluconazole (41), we were interested in analyzing the role of CaFZO1 in azole tolerance in C. albicans. We tested the tolerance of fzo1Δ/Δ mutant to azoles and polyenes (drugs that target the membrane) by spot assays and MIC determination. The fzo1Δ/Δ mutant was unable to grow in the presence of the azole antifungals tested, whereas it was moderately susceptible to the polyene, amphotericin B, indicating that dysfunctional mitochondria may affect susceptibility to azoles (Fig. 4A). The MIC₈₀ value for the fzo1Δ/Δ mutant in response to fluconazole was reduced by a modest 4-fold and 2-fold for amphotericin B, whereas it was significantly reduced by 7.5-fold for itraconazole and ketoconazole compared to the wild type and reconstituted strain (Table 4).

Table 4.

Antifungal susceptibilities of C. albicans fzo1Δ/Δ mutants

Antifungal	MIC80 (μg ml−1)			Fold change
	Wild type	fzo1Δ/Δ	fzo1Δ/Δ+FZO1
Fluconazole	0.5	0.125	0.5	4
Itraconazole	0.015	0.002	0.078	7.5
Ketoconazole	0.015	0.002	0.015	7.5
Amphotericin B	0.625	0.3125	0.625	2

The role of drug efflux pumps, CDR1 and CDR2 (ABC transporters), and MDR1 (MFS transporter) in tolerance to azole antifungals is well documented in C. albicans (42). Since the susceptibility to azole antifungals was affected in the fzo1Δ/Δ mutant, we were prompted to analyze the status of CDR1 in the mutant. Genome-wide transcriptome data revealed that there was no change in the transcript levels of CDR1 and CDR2 compared to the wild type (Fig. 4B). On the contrary, transcript levels of MDR1 were significantly higher in the mutant, as confirmed by qPCR (Fig. 4B). This is consistent with a previous report, wherein upregulation of MDR1 in a petite mutant strain of C. albicans is documented (Fig. 4B) (8). Given that mitochondria are the major source of ATP and the efflux activity of CDR pumps is energy dependent, the ability to efflux rhodamine 6G (R6G), a substrate of CDR pumps, was analyzed in the mutant (43). No efflux of R6G was observed without the addition of glucose in all of the strains. Upon the addition of glucose, an increase in the extracellular concentration of R6G was observed from 0.59 to 4.56 nmol ml⁻¹ in 10 min (7.7-fold increase) in the wild type (Fig. 4C). Similarly, a 6.3-fold increase was observed in the reconstituted strain. In contrast, the fzo1Δ/Δ mutant exhibited a 2.2-fold increase in extracellular R6G, indicating a deficiency in R6G transport, probably due to the reduced activity of CDR1 (Fig. 4C).

Cdr1p regulation can occur at transcriptional, posttranscriptional, and posttranslational levels (44). The decreased activity of CDR1 could also be attributed to low protein levels on the plasma membrane (PM) due to its missorting in the mutant. In order to assess the cause for low activity, we used a translational CDR1-GFP reporter fusion construct (45), which was integrated at its native locus in the wild-type and mutant genomes. Therefore, the expression of the CDR1-GFP fusion protein was driven from the CDR1 promoter. The CDR1-GFP fluorescence in actively growing cells (OD₆₀₀ = 1.0) was visualized by confocal microscopy and quantified by flow cytometry (Fig. 4D). Interestingly, in the mutant, CDR1-GFP was distributed between the PM and the vacuole compared to its PM localization in the wild type (Fig. 4D). Moreover, the intensity of the PM localized Cdr1p was slightly lower in the mutant compared to the wild type. The presence of CDR1-GFP in the vacuole is suggestive of its missorting in the mutant and also points to the possibility of its increased degradation and/or reduced half-life.

Fluphenazine, a documented inducer of CDR1 expression, was used in order to test the inducibility of CDR1-GFP in wild-type and mutant backgrounds (46). The expression of CDR1-GFP was responsive to fluphenazine in both strains, as indicated by significantly high fluorescence intensity on the PM, compared to the uninduced condition (Fig. 4D). Interestingly, the fold induction between the wild type and the mutant remained the same (∼2-fold), arguing against transcriptional control, in line with the qPCR data (Fig. 4D). We observed that whereas CDR1-GFP was distributed between the PM and the vacuole, the PM localization of Cdr1p was decreased in the mutant, similar to the uninduced condition. The difference in the total fluorescence intensity, as obtained by flow cytometry analysis, between the wild type and the mutant was reflected better in the induced condition than in the uninduced condition. The total fluorescence intensity was significantly decreased in the mutant (16.8 ± 0.8) compared to the wild type (21.9 ± 1), a finding suggestive of lower protein levels in the mutant (Fig. 4D). Taken together, the data indicate that the absence of functional mitochondria leads to decreased surface delivery of Cdr1p due to its mistargeting to the vacuole.

Absence of FZO1 results in increased expression of iron uptake genes.

Iron forms an essential component in metalloproteins such as cytochromes and Fe-S proteins involved in respiration, DNA synthesis, oxygen storage, and essential metabolic pathways in C. albicans. Moreover, iron availability in C. albicans has been linked to drug resistance and morphology of this fungus (47, 48). In the present study, we also link dysfunctional mitochondria to iron homeostasis in C. albicans. Transcriptome analysis of fzo1Δ/Δ cells showed transcriptional changes in genes involved in iron homeostasis. A total of 27 genes associated with iron assimilation/transport were up (n = 13)- or down (n = 14)-regulated in the mutant; 7 genes of these genes matched those that are induced in iron starvation conditions in C. albicans. Reductases and oxidoreductases involved in iron uptake (FRE2, FRE5, and FRE31), siderophore transporter (ARN1/SIT1), copper transporter ATPase (CCC2), heme-utilizing cell wall protein (RBT5), and a transcriptional activator of respiration (HAP32) were highly upregulated in fzo1Δ/Δ mutant (Fig. 3C and 5A). Homologs of these genes in S. cerevisiae are known to be induced in low-iron conditions (49, 50), wherein ScARN1, ScFRE3, and ScCCC2 are also upregulated in ρ⁰ cells (51).

Fig 5.

(A) qPCR of selected genes from transcriptome analysis involved in iron homeostasis. The fold change was calculated by using 2^−ΔΔCT, normalized to ACT1 (endogenous control), with the wild type as the calibrator either in basal Fe³⁺ or in supplemented Fe³⁺ (100 μM) conditions. Values are means ± the SD and are derived from two independent RNA preparations. (B) Fivefold serial dilutions of cell suspensions were spotted on indicated medium plates supplemented with FeSO₄ at the indicated concentrations, followed by incubation at 37°C for 48 h. WT, wild type.

To test the hypothesis that mitochondrial-function-challenged fzo1Δ/Δ cells mimic iron-starved conditions and upregulation of the iron uptake genes was responsive to iron levels, exogenous Fe³⁺ was supplemented in the media. Wild-type and fzo1Δ/Δ cells were cultured in 100 μM FeCl₃ and expression of the above genes was measured by qPCR. Wild-type transcript levels of the mentioned genes were unaffected with addition of exogenous FeCl₃. Although ARN1 was upregulated (73.7 ± 2.7)-fold (in the mutant) in basal Fe³⁺ levels (the medium contains 1.23 μM Fe³⁺), FeCl₃ supplementation reduced the fold change to 3.1 ± 0.8. Similarly, the transcript levels of FRE31, FRE2, FRE5, RBT5, CCC2, and HAP32 were found to be iron responsive solely in the fzo1Δ/Δ mutant (Fig. 5A), reaffirming deregulation of the iron metabolism in the mutant.

In S. cerevisiae, loss of mtDNA leads to generation of the iron starvation response as a result of altered mitochondrial Fe-S cluster (ISC) biogenesis or its export into the cytosol (51–53). Since the transcriptome analysis was performed in iron-replete conditions and considering that the fzo1Δ/Δ cells were devoid of mtDNA, we hypothesized that the generation of the iron-starved response in the fzo1Δ/Δ cells could be a result of perturbed Fe-S cluster biogenesis or the export of the Fe-S cofactors into the cytosol. One of the consequences of an impaired ISC biogenesis is the activation of the iron regulon causing increased iron uptake and hence increased intracellular free iron levels, which cause oxidative stress (53–55). Therefore, as an indirect indicator of perturbed Fe-S biogenesis, we tested fzo1Δ/Δ cells for growth sensitivity to FeSO₄. The demand for iron is high when cells are grown in respiratory (glycerol) and respirofermentative (galactose) media since they need Fe-containing proteins for oxidative phosphorylation. Hence, the mutant was tested for growth on all of the aforementioned media supplemented with FeSO₄ incubated at 30 and 37°C. We observed that, irrespective of the medium used, there was no difference in the growth of the mutant in the presence or absence of FeSO₄ (Fig. 5B). FeSO₄ concentrations as high as 160 mM in glycerol- or galactose-containing medium did not hinder the growth of the mutant relative to the wild type (data not shown). Nevertheless, considered together, our data suggest that dysfunctional mitochondria in fzo1Δ/Δ cells cause a defect in iron homeostasis.

Oxidative stress induced activation of the HOG1 pathway is impaired in fzo1Δ/Δ cells.

An increase in ROS occurs in fungal cells in response to oxidative stress (56). Adapting to stress conditions such as treatment with peroxide or high salinity requires energy production and hence mitochondrial activity (57, 58). Altered mitochondrial iron levels also lead to hypersensitivity to oxidative stress, which is reflected in sensitivity to ROS-inducing agents such as H₂O₂ (54, 59, 60). Therefore, we tested the growth of fzo1Δ/Δ cells in the presence of H₂O₂ in fermentative (glucose) and respirofermentative (galactose) media. Whereas H₂O₂ moderately inhibits growth of the mutant in glucose containing media, the growth defect is pronounced when galactose was provided as the carbon source (Fig. 6A).

Fig 6.

(A) Fivefold dilutions of strains were spotted onto YNB-glucose or YNB-galactose plates containing the indicated concentrations of H₂O₂ and grown for 48 h at 30°C. (B) Hog1p is not phosphorylated in fzo1Δ/Δ cells in the presence of H₂O₂. Western blotting of protein extracts from the indicated strains was performed after a 10-min treatment with 10 mM H₂O₂. (C) Strains were grown in YNB-galactose (2%) till early log phase, washed twice in PBS, and incubated in 10 μM DCFDA for 30 min for intracellular ROS measurement. Wild-type cells were treated with 10 mM H₂O₂ for 30 min prior to staining as a positive control. The values indicate mean percentages ± the SD (n = 3) of stained cells. *, P < 0.01 (Student t test). (D) qPCR of AOX2 in mutant cells, where the relative expression was calculated by using 2^−ΔΔCT, normalized to ACT1 (endogenous control), with the wild type as the calibrator. WT, wild type.

Two MAPKs, Hog1 and Mkc1, are activated upon oxidative stress (56, 61). The phosphorylation of Mkc1 is dependent on the HOG pathway; therefore, the latter is considered the major oxidative stress-responsive pathway (61, 62). Cells growing in exponential phase were challenged with 10 mM H₂O₂, and their protein extracts were analyzed by Western blotting. As expected, the Hog1 and Mkc1 pathways were not activated in the fzo1Δ/Δ mutant upon exposure to 10 mM H₂O₂ for 10 min, whereas the wild-type and reconstituted strains showed sufficient activation (Fig. 6B). The absence of P-Mkc1 protein in the mutant is in line with an earlier report where, upon oxidative stress, Mkc1 activation was shown to be dependent on Hog1 (61).

Since adaptation to oxidative stress is impaired in fzo1Δ/Δ cells and mitochondrial dysfunction is associated with increased ROS levels, we hypothesized that an impaired HOG cascade will result in increased cellular ROS. Cellular ROS measured by DCFDA showed a 2-fold increase in the fzo1Δ/Δ mutant comparable to wild-type cells treated with 10 mM H₂O₂ for 30 min (Fig. 6C). Induction of the alternate respiratory (AOX) pathway as a defense mechanism against oxidative stress has been documented in plants (63). In C. albicans, this cyanide-insensitive pathway, consisting of the constitutive AOX1a and the oxidant inducible AOX1b (AOX2), is induced by aging and stresses mediated by H₂O₂, menadione, and paraquat (27). Consistent with these facts, AOX2 was the most upregulated gene in transcriptome profiling of fzo1Δ/Δ cells, a finding also validated by qPCR (Fig. 6D). These results suggest that altered mitochondrial function leads to increased susceptibility to H₂O₂ via an impaired HOG pathway and high cellular ROS, concomitant with elevated AOX2 transcript. Taken together, these results link mitochondria to the activation of the oxidative-stress-induced Hog1 pathway.

Mitochondrial phospholipid and cellular ergosterol levels are altered in the mutant.

The endoplasmic reticulum (ER) is the major site for the biosynthesis of all phospholipids, except for phosphatidylethanolamine (PE) and cardiolipin (CL), which are synthesized in the mitochondria. The transport of phosphatidylserine (PS) from the ER to the mitochondrion and its subsequent conversion to PE, which is transported back to the ER for phosphatidylcholine (PC) synthesis, involves mechanistically unresolved trafficking steps between the two organelles (Fig. 7A). The ER-mitochondrion encounter structure complex (ERMES), Mmm1-Mdm10-Mdm12-Mdm34, physically connects the ER to the mitochondria and is involved in mitochondrial morphology and genome maintenance (64, 65). Moreover, studies in C. albicans show an association between phospholipid homeostasis and mitochondrial function (41, 66).

Fig 7.

(A) Simplified schematic of phospholipid trafficking in yeast cells. (B) The indicated yeast cells were cultivated in YNB-glucose in the presence of ³²P_i. Phospholipids were extracted from crude mitochondrial fractions and separated by TLC. Steady-state phospholipid species were determined. (C) The amounts of each lipid relative to total phospholipids were determined and are presented as percentages. fzo1Δ/Δ-1 and fzo1Δ/Δ-2 are two independent null mutants. Values are means ± the SD (n = 6). *, P < 0.01 (Student t test). CL, cardiolipin; PA, phosphatidic acid; PE, phosphatidylethanolamine; PS, phosphatidylserine; PI, phosphatidylinositol; PC, phosphatidylcholine; PG, phosphatidylglycerol; CDP-DAG, cytidine diphosphate diacylglycerol; LPC, lysophosphatidylcholine; LPI, lysophosphatidylinositol.

We therefore analyzed fzo1Δ/Δ cells for defects in the steady-state levels of phospholipids by growing wild-type, mutant, and reconstituted strains in ³²P_i-containing media. Total phospholipids were extracted from the crude mitochondrial membranes and separated by TLC (Fig. 7B). Although the overall differences in phospholipid content between the wild-type and mutant strains were modest, the mutant displayed an increased amount of PE compared to the wild-type (Fig. 7C). The amount of PS also increased, whereas the amounts of PC, PG (phosphatidylglycerol), and CL decreased in the mutant, indicating that either the rate of conversion of PS to PE and PC was affected due to the altered activity of Psd1p (phosphatidylserine decarboxylase 1) or there was reduced lipid trafficking between the ER and the mitochondria (Fig. 7C). The decreased levels of PG and CL were small but significant. Studies have shown that there are high PE levels in CL-deficient mutants and high CL levels in PE-deficient mutants in S. cerevisiae (67, 68). Therefore, high PE levels could be a compensatory response to low CL levels in fzo1Δ/Δ cells. Alternatively, given that the decrease in CL is relatively modest, the increased abundance of PE could instead be explained by an increased activity of Psd1 in the mutant, similar to what has been shown in sam37Δ/Δ cells (10). Considering that mitochondrial membranes are enriched in PC (69), another possibility that could partly account for the decreased levels of PC could be the defect in mitochondrial morphology in fzo1Δ/Δ cells. Moreover, the loss of ERMES subunits, Ups1p, or Mdm31p leads to similar phospholipid profiles such as decreased CL levels and increased PS levels in S. cerevisiae (70). This indicates that deletion of FZO1 in C. albicans affects phospholipid levels. Consistent with a perturbed iron homeostasis in the mutant, the transcriptome data revealed downregulation of five genes involved in the ergosterol biosynthesis pathway (Fig. 8A): ERG1 (squalene epioxidase), ERG3 (C-5 sterol desaturase), ERG4 (protein similar to sterol C-24 reductase), ERG11 (lanosterol 14α demethylase), and ERG25 (putative C-4 methyl sterol oxidase) (Table 3). The downregulation of these genes was also validated by qPCR (Fig. 8B). Since the ergosterol biosynthetic pathway is regulated by iron availability in yeast, the altered expression pattern of these genes in the mutant may be an indirect response to a deregulated iron homeostasis. In S. cerevisiae, perturbation of the ISC systems also leads to repressed respiration and heme metabolism (52). In lieu of these facts, it is possible that ERG11, a cytochrome P-450-dependent enzyme of the ergosterol biosynthesis pathway, was downregulated in the mutant since it requires heme for its activity (37). In order to correlate the reduced transcript levels of the ERG genes to total ergosterol levels, we subjected the fzo1Δ/Δ mutant to GC-MS analysis. The analysis revealed a decrease in cellular ergosterol content compared to the wild type. The mutant had the lowest ergosterol content at 2.43 mg g of lipid⁻¹ compared to 144.5 and 127.9 mg g of lipid⁻¹ in the wild type and revertant, respectively (Fig. 8C). A decrease in ergosterol levels is in line with downregulated transcript levels of ERG11 and other ERG genes (ERG3 and ERG1). These data therefore indicate that the mitochondrial status can influence the ergosterol biosynthetic pathway as an indirect consequence of altered iron homeostasis.

Fig 8.

(A) Schematic representation of the ergosterol biosynthetic pathway in C. albicans. The genes indicated in boldface were downregulated in fzo1Δ/Δ cells. (B) qPCR analysis of ERG1, ERG3, and ERG11, normalized to ACT1, where the average relative expression (calculated by 2^−ΔΔCT) is derived from two independent RNA preparations, and the SD is indicated by error bars. (C) Ergosterol content of strains. Ergosterol was extracted from cells as described and quantified by GC-MS analysis. Values are described as ergosterol (mg g of lipid⁻¹). All values are means ± the SD of four independent assays. *, P < 0.05 (Student t test).

The susceptibility of an azole-resistant clinical isolate to azole antifungals is affected upon deletion of FZO1.

To investigate the contribution of mitochondria to the development of drug resistance in clinical isolates, we deleted (SAT-flipper strategy) FZO1 in two well-studied matched pairs of azole-susceptible (GU4) and azole-resistant (GU5) isolates (71). Deletions were confirmed by Southern hybridization (data not shown). MGFM and DAPI staining confirmed the presence of aberrant mitochondria and the loss of mtDNA in both GU4 and GU5 (data not shown). The GU4 fzo1Δ/Δ cells displayed increased susceptibility to fluconazole, ketoconazole, and itraconazole antifungals as reflected in the MIC₈₀ values (Table 5). The MIC₈₀ value for GU4 fzo1Δ/Δ was reduced by 4-fold for fluconazole and ketoconazole and by 8-fold for itraconazole. Similarly, for the GU5 fzo1Δ/Δ cells, the MIC₈₀ value was reduced by a modest 2-fold and 4-fold for fluconazole and ketoconazole, respectively, and by 8-fold for itraconazole (Table 5). This data set demonstrates that the susceptibility of azole-resistant clinical isolates to azole antifungals can be marginally increased by targeting mitochondrial functions.

Table 5.

Antifungal susceptibilities of C. albicans fzo1Δ/Δ mutants in a matched pair of clinical isolates

Antifungal	GU4				GU5
	MIC80 (μg ml−1)			Fold change	MIC80 (μg ml−1)			Fold change
	Wild type	fzo1Δ/Δ	fzo1Δ/Δ+FZO1		Wild type	fzo1Δ/Δ	fzo1Δ/Δ+FZO1
Fluconazole	3.125	0.78125	1.562	4	>100	50	100	2
Ketoconazole	0.0243	0.00609	0.04875	4	0.04875	0.0121	0.0243	4
Itraconazole	0.0975	0.0121	0.04875	8	0.04875	0.00609	0.0243	8

DISCUSSION

The identification of mitochondria as a determinant of drug susceptibility, cell wall damage, phospholipid homeostasis, and virulence has led to a considerable focus on this organelle in C. albicans (72). The basis of our study is a mutant, which has been created by deleting CaFZO1, a component of the yeast mitochondrial fusion machinery, thereby directly targeting the maintenance of functional mitochondria (14). Our study provides new insight into the molecular basis of azole susceptibility and oxidative stress, phenotypes that are linked to the presence of functional mitochondria. Furthermore, we extend the role of mitochondria to iron homeostasis and link it to cellular processes like ergosterol biosynthesis. fzo1Δ/Δ cells display fragmented mitochondria, loss of mitochondrial genome, and reduced mitochondrial membrane potential. Growth defects of the mutant (Fig. 1C) and its inability to grow on a nonfermentable carbon source (glycerol + ethanol) (Fig. 2A) translates altered mitochondrial morphology to loss of its function. Loss of mtDNA as a consequence of altered mitochondrial morphology was also validated (Fig. 2C), further confirming the contribution of FZO1 in maintaining mitochondrial morphology and subsequently cell metabolism.

Previous studies with S. cerevisiae and C. glabrata indicate that loss of mitochondrial function leads to an elevated expression of the ABC transporter genes (ScPDR5, CgCDR1, and CgCDR2) via transcription factors ScPdr3 and CgPdr1, causing the multidrug resistance (MDR) phenotype (39, 40, 73, 74). Here, we show that in C. albicans, whereas aberrant mitochondria, modestly affects susceptibility to fluconazole, there is a significant reduction in MIC₈₀ values for ketoconazole and itraconazole (Table 4). On the contrary, the growth of the fzo1Δ/Δ cells was abrogated in the presence of azoles in spot assays (Fig. 4A). Given the slow-growth phenotype of the mutant in the absence of azoles, the abrogated growth of the mutant in spot assays in the presence of azoles may be a result of inadequate ATP supply or the arrest of energy-dependent cellular processes due to dysfunctional mitochondria. Nevertheless, we attempted to understand the molecular basis of azole susceptibility by addressing the status of the ATP-dependent major efflux pump, CDR1, in the mutant. Only one other study in C. albicans has linked a mitochondrial localized protein, Goa1p, with azole sensitivity and abrogated efflux activity of Cdr1p, along with a commensurate decrease in its transcript levels (12). In the present study, we show that the CDR efflux pump activity is significantly reduced but not abrogated. Localization studies on CDR1 revealed that the protein is largely missorted to the vacuole in the mutant, indicating that functional mitochondria exerts posttranslational regulation on Cdr1p levels that could reduce the stability/half-life of the efflux pump protein. As a consequence, there is reduced protein on the PM, which could partly contribute to the decreased efflux activity of Cdr1p (Fig. 4C and D). Elsewhere, it has been shown that reduced PM localization of Cdr1p as a result of its missorting is associated with increased susceptibility to azoles (45, 75). Our data rule out transcriptional regulation of CDR1 based on two observations: (i) no change in the transcript levels of CDR1 and CDR2 was observed in the mutant compared to the wild type and (ii) fluphenazine-induced fold induction of CDR1-GFP remained the same (∼2-fold) between the wild type and the mutant (Fig. 4B and D). We conclude that the reduced activity of Cdr1p in cells with dysfunctional mitochondria may likely extend beyond its transcriptional regulation in C. albicans. As a consequence, susceptibility to azoles is affected in fzo1Δ/Δ cells, albeit to modest levels, indicating that mitochondrial function may have a limited role in azole susceptibility. Another implication of this result is that not all means of eliminating mitochondria in C. albicans will have similar consequence with respect to azole susceptibility.

Transcriptome profiling of the mutant revealed “downregulation” as the major consequence to the loss of mitochondria (Fig. 3A and B). This implies that the loss of mitochondria elicits a large-scale cellular response involving a major shutdown of genes related to mitochondrial function, transporters, lipid metabolism, and iron homeostasis. Furthermore, in order to provide an understanding into the cellular behavior vis-à-vis cellular adaptability in the mutant, we used network modeling to define sets of genes that are coregulated (Fig. 3C). Mitochondrial biogenesis is tightly controlled and is coordinated with cell division cycle. Consistent with this observation, there was a subset of downregulated genes associated with cell cycle. Previous studies with Goa1p and Sam37p demonstrate a link between mitochondria and cell wall justifying the downregulation of cell wall organization genes, a finding in line with the transcriptome profiling of goa1Δ/Δ cells (12). Most striking of the coregulated subsets was the one that contained genes associated with iron homeostasis, the implications of which are discussed below. Gene coregulatory network analysis therefore allowed us to gain further insight into functional modules that are induced in response to dysfunctional mitochondria. Whether the network structure will be carried forward onto protein-protein interaction network needs to be investigated.

Comprehensive analysis of transcriptome data revealed derepression of a cluster of genes associated with iron uptake, namely, RBT5, SIT1, CCC2, FRE31, FRE2, and FRE5 (Table 3). These genes are known to be upregulated in iron-restrictive conditions (36). The expression of these genes was iron dependent, as is evident from a decrease in their transcript levels upon iron supplementation (Fig. 5A). In S. cerevisiae, the loss of mtDNA leads to a signature transcriptional response of iron starvation. In addition, mitochondria are the major site for ISC biogenesis, and the disruption of the ISC assembly and export machineries induces the accumulation of iron within mitochondria (76), leading to the constitutive expression of iron-regulated genes (77, 78). Expression profiles of cells that are iron starved largely overlap with those of cells that are defective in mitochondrial ISC biogenesis (52). A recent study (53) in S. cerevisiae shows that cardiolipin deficiency leads to altered mitochondrial and cellular iron homeostasis, causing an increased expression of the iron regulon genes, elevated mitochondrial iron levels, and susceptibility to ROS-inducing agents. All of these phenotypes are consequences of perturbed ISC biogenesis in the mutant (53). Considering that our transcription profiling was performed in iron-replete conditions, we proposed that increased expression of the iron uptake genes could be due to perturbed ISC biogenesis, which would lead to elevated mitochondrial iron levels in the mutant. Testing the fzo1Δ/Δ cells for iron sensitivity, as an indirect measurement for increased cellular iron levels, showed that, unlike S. cerevisiae, the mutant was able to tolerate increasing concentrations of iron. Therefore, based on iron sensitivity assay, we were unable to link deregulated iron homeostasis to perturbed ISC biogenesis in the present study. The existence of additional mechanisms that could enable the mutant to tolerate high concentrations of iron cannot be ruled out, based on our result. Studies in C. albicans linking mitochondria to iron homeostasis are limited. Mrs4, a mitochondrial carrier protein required for mitochondrial morphology is involved in the maintenance of cellular iron content. The mrs4Δ/Δ cells display increased cellular iron content and altered expression of iron regulon genes. In addition, reduced expression of CaSMF3 (putative vacuolar iron transporter) in the mrs4Δ/Δ cells affects the vacuolar transport of iron to the cytosol, also affecting the cellular iron content (79). In view of these facts, it can be proposed that deregulated iron homeostasis could be a common underlying effect in cells with dysfunctional mitochondria. Whether the altered iron homeostasis in fzo1Δ/Δ cells is actually due to perturbed ISC biogenesis, as suggested from studies in S. cerevisiae, needs to be investigated. In this context, direct measurement of intracellular iron levels in the mutant will aid in correlating the increased expression of the iron uptake genes to altered ISC biogenesis in the fzo1Δ/Δ cells. Recently, involvement of Hog1p, the conserved stress activated MAP kinase, in response to iron availability has been demonstrated in C. albicans. The hog1Δ/Δ cells display increased expression of several iron uptake genes under iron-replete conditions, similar to fzo1Δ/Δ cells in the present study (80). In view of these facts, deregulated iron homoeostasis in fzo1Δ/Δ cells could also be an indirect effect of the impaired activation of the HOG pathway (Fig. 6B). The finding that numerous cellular processes, such as heme metabolism and ergosterol biosynthesis, linked to iron metabolism are affected in fzo1Δ/Δ cells (Table 3, Fig. 5A and 7) strengthens the connection between functional mitochondria and iron homeostasis. Together, our data add another dimension to the profound effects of mitochondrial dysfunction in C. albicans.

Mitochondria in general is known to counter effects due to increased ROS production, and the regulatory role of HOG pathway in response to oxidative stress is also known (56). A link between Hog1p and respiratory metabolism has also been demonstrated in C. albicans, wherein the deletion of HOG1 leads to an increased dependence on mitochondrial ATP synthesis (26). Consistent with these facts, the fzo1Δ/Δ cells were sensitive to H₂O₂ and displayed increased ROS production. Interestingly, fzo1Δ/Δ cells were also unable to activate the HOG pathway, upon peroxide stress (Fig. 6A, B, and C). These results, for the first time, link peroxide-induced activation of Hog1p to the presence of functional mitochondria, implying a key role for mitochondria in controlling this signaling pathway. In aggregate, our results suggest a connection between functional mitochondria, activation of HOG pathway, and iron homeostasis in C. albicans. During oxidative stress, the transcription factor (TF) Cap1 is activated which, along with the Mrr1 TF, induces MDR1 expression (81). Therefore, the observed increased expression of MDR1 in fzo1Δ/Δ cells (Fig. 4) could be explained by the prevailing oxidative stress condition in the mutant.

Genes associated with ergosterol biosynthetic pathway are regulated by iron availability in yeast (82). In agreement with the deregulated cellular iron homeostasis in the mutant, five genes in the ergosterol biosynthesis pathway were downregulated, with a commensurate decrease in ergosterol levels (Fig. 8B and C). In addition, low levels of heme may also contribute to the downregulation of the ergosterol biosynthesis pathway, since ERG11 is a heme-containing enzyme (37). Of the genes involved in heme biosynthesis, HEM14 (putative protoporphyrinogen oxidase) and HEM13 (coproporphyrinogen III oxidase) were significantly downregulated (Table 3). Therefore, our study extends the link between mitochondria and deregulated iron homeostasis to an effect on ergosterol biosynthesis pathway. Ergosterol and sphingolipids are considered important determinants of PM localization of Cdr1p. Mutants defective in either of the pathways display reduce PM localization and reduced efflux activity of Cdr1p, leading to increased susceptibility to azoles (45, 83). Considering these facts and taking into account that Cdr1p was missorted in the mutant, we deduce that low ergosterol may cause membrane perturbation, thereby leading to the reduced activity of Cdr1p. Therefore, we interpret that the altered membrane environment could also be the basis for the reduced activity of Cdr1p in the absence of functional mitochondria.

Compensatory changes on the cell membrane occur in situations where the levels of one constituent of the membrane are altered (low ergosterol level in the mutant). A connection between Sam37p and PE biosynthesis has been demonstrated in C. albicans, wherein sam37Δ/Δ cells have high Psd1 activity but are compromised in PE biosynthesis because of low conversion rates of PS to PE, indicating that Sam37 functions to traffic phospholipids between the ER and the mitochondria (41). Analysis of steady-state levels of mitochondrial phospholipids in the fzo1Δ/Δ cells showed high levels of PE and low levels of PC (Fig. 7B and C), a finding suggestive of either altered activity of Psd1 or disruption of the ERMES complex leading to trafficking defects. Given the morphological defect in our mutant and that phospholipid trafficking is important for phospholipid metabolism, altered levels of phospholipids could be a consequence of trafficking defects. However, the overall differences in phospholipid content between the wild-type and mutant strains were modest. Considering that the maintenance of specific phospholipid composition in the outer and inner mitochondrial membranes is crucial for vital mitochondrion-related functions, we speculate that there may be an activation of compensatory mechanisms in the mutant to maintain phospholipid homeostasis (84, 85). As a result, the mutant displays only a minor change in the percentage of total phospholipids. Whether this modest but significant alteration in phospholipid levels in fzo1Δ/Δ cells contributes to the effects observed therein remains to be addressed. These results nonetheless indicate the importance of mitochondrial fusion and fission in maintaining a connection between the mitochondrion and the ER for phospholipid homeostasis.

Keeping in view the pleiotropic effects displayed by cells with dysfunctional mitochondria, we surmise that targeting mitochondrial functions would have therapeutic implications in C. albicans. Of the two proteins used to target mitochondrial functions in C. albicans, Goa1p is unique to the CTG clade of the subphylum Saccharomycotina, which excludes C. glabrata. Sam37, on the other hand, has orthologs in humans, but Pfam-based characterization of the domain structures suggests differences in the structure and function of the fungal and human proteins (10). CaFZO1 is unique to the fungal kingdom, with strong orthologs in the genomes of Saccharomyces cerevisiae, Schizosaccharomyces pombe, Magnaporthe grisea, Aspergillus niger, and Neurospora crassa but with no known human or murine ortholog (17), an observation suggestive of its potential to be used as a target for developing new antifungals. However, the fact that impairment of mitochondrial function in C. glabrata results in increased resistance to azoles precludes this approach of mitochondrial inhibition as universally suitable for all pathogenic yeast and emphasizes the importance of revealing species specific mechanisms of action. The use of azole antifungals in combination with mitochondrial inhibitors would increase their efficacy and may prove useful for reversing the susceptibility of azole-resistant isolates in C. albicans. To date, an association of dysfunctional mitochondria to azole tolerance has been demonstrated in the clinical isolates of the pathogenic fungi such as C. glabrata, C. albicans, and Cryptococcus sp. (7, 86–88). In this regard, our studies in GU4 and GU5 are relevant since these strains are serial isolates obtained from an AIDS patient with recurrent episodes of oropharyngeal candidiasis (71). Deletion of FZO1 rendered cells modestly sensitive to azoles, except for itraconazole in both GU4 and GU5, indicating a minor role for mitochondria in azole resistance in this pair of clinical isolates (Table 5). Nevertheless, targeting mitochondria in various clinical isolates will provide insight into the contribution of this organelle to the development of drug resistance and will highlight additional molecular mechanisms operating in these strains. Therefore, our study adds to the list of proteins that may be used as a target to intervene with mitochondrial functions in C. albicans.