Multifactor transcriptional control of alternative oxidase induction integrates diverse environmental inputs to enable fungal virulence

Liu, Zhongle; Basso, Pauline; Hossain, Saif; Liston, Sean D.; Robbins, Nicole; Whitesell, Luke; Noble, Suzanne M.; Cowen, Leah E.

doi:10.1038/s41467-023-40209-w

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Published: 27 July 2023

Multifactor transcriptional control of alternative oxidase induction integrates diverse environmental inputs to enable fungal virulence

Nature Communications volume 14, Article number: 4528 (2023) Cite this article

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Abstract

Metabolic flexibility enables fungi to invade challenging host environments. In Candida albicans, a common cause of life-threatening infections in humans, an important contributor to flexibility is alternative oxidase (Aox) activity. Dramatic induction of this activity occurs under respiratory-stress conditions, which impair the classical electron transport chain (ETC). Here, we show that deletion of the inducible AOX2 gene cripples C. albicans virulence in mice by increasing immune recognition. To investigate further, we examined transcriptional regulation of AOX2 in molecular detail under host-relevant, ETC-inhibitory conditions. We found that multiple transcription factors, including Rtg1/Rtg3, Cwt1/Zcf11, and Zcf2, bind and regulate the AOX2 promoter, conferring thousand-fold levels of inducibility to AOX2 in response to distinct environmental stressors. Further dissection of this complex promoter revealed how integration of stimuli ranging from reactive species of oxygen, nitrogen, and sulfur to reduced copper availability is achieved at the transcriptional level to regulate AOX2 induction and enable pathogenesis.

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Introduction

Fungi have a profound, but underappreciated impact on human health, killing more than 1.5 million of the billions of people they infect worldwide each year^1,2,3,4. Vulnerability to fungal infection is greatest for individuals with impaired immune function⁵. Of the deaths attributable to fungal infection in North America, ~90% are caused by species of Candida, Aspergillus, and Cryptococcus⁶. Among these, Candida albicans is one of the most frequent causes of invasive infection, with systemic candidiasis causing considerable morbidity and driving mortality rates as high as 40% despite treatment⁷. Poor outcome for invasive infections is attributable to the limited number of effective antifungals available and an alarming rise in frequency of clinical resistance.

The ability to utilize alternative carbon sources plays an essential role in allowing C. albicans to transition from a commensal colonizer to a pathogenic microbe^8,9,10. During infection, this metabolic flexibility contributes to virulence and the emergence of drug resistance¹¹. In particular, pathogenic fungi rely on respiration to utilize the poorly-fermentable carbon sources in host environments, not only for energy generation, but also to support essential anabolic processes^12,13.

Under conditions compromising the classical electron transport chain (ETC), C. albicans makes use of a bypass respiratory pathway, which is supported by cyanide-insensitive alternative oxidases (Aox) located on the matrix side of the inner mitochondrial membrane^14,15. Encoded by nuclear genes, constitutive (Aox1) and highly inducible (Aox2) isoforms catalyze electron transfer from ubiquinol to oxygen, without coupling it to proton-pumping. Found in diverse microbial pathogens, but not in humans, Aox is proposed as a potential drug target given its reported role in governing diverse cellular stress response pathways, including a role in maintaining cell wall architecture^16,17. Early-stage studies with inhibitors, either alone or in combination with inhibitors of the ETC, have shown promise^16,18,19. In C. albicans and other fungi, the importance of the classical ETC for virulence is well established^16,20,21. In contrast, how Aox activity is regulated in response to stressors encountered during infection and how induction contributes to virulence have remained enigmatic.

Results

Inducible Aox activity supports systemic virulence in mice

Candida spp. and other pathogenic microbes possess a bypass respiratory pathway supported by cyanide-insensitive alternative oxidases (Aox), which are located on the matrix side of the inner mitochondrial membrane (Fig. 1a). To learn whether Aox activity plays a role in the virulence of C. albicans, we focused primarily on AOX2 due to its high level of inducibility in response to environmental challenges. We constructed an AOX2 homozygous deletion mutant (referred to as aox2-deletion or aox2; the same nomenclature is used to reference all homozygous deletion mutants throughout this study). Upon systemic infection of immunocompetent mice, the aox2-deletion strain was significantly less virulent than the parental strain (Fig. 1b). Restoration of either allele of AOX2(A) or AOX2(B) was sufficient to restore virulence to a level equivalent to that of the wild-type strain (Fig. 1b).

**Fig. 1: *AOX2* induction enables virulence in mice.**

To assess the proliferative capacity of the aox2-deletion strain within an infected host, we measured fungal burden in major organs following three days of systemic infection. The numbers of colony-forming units (CFU) recovered from aox2-infected kidneys, livers, spleens, and brains were comparable to those from tissues infected by the complemented strain (Fig. 1c). This result indicated that, although AOX2 is required for progressive, lethal infection, it is dispensable for early events in pathogenesis. To determine whether differences in the host response contribute to the contrasting outcomes of infection with aox2-deletion vs. the complemented strain, we examined the extent of immune-cell accumulation in infected kidneys (organ of greatest fungal burden). Representative fixed sections of mock-infected, aox2-infected, and aox2 + AOX2(A)-infected renal cortex are shown in Fig. 1d; as indicated in the figure panels, fungal cells were stained with anti-Candida antibody and immune cells with antibody against the leukocyte common antigen (LCA, also known as CD45). Throughout the renal cortex, invading aox2 fungi induced markedly greater leukocyte accumulation than the complemented strain (Fig. 1d). Large dense foci of CD45⁺-cells were observed in non-contiguous sections of kidneys recovered from independent mice infected with the aox2-deletion strain. In contrast, relatively sparse, diffuse infiltration was observed in kidneys infected by the aox2 + AOX2(A) strain (Fig. 1d; Supplementary Fig. 1a). Thus, greater leukocyte accumulation could be observed in the kidney during challenge with the aox2 strain than with the complemented strain, even though overall fungal burdens were comparable at this very early stage of infection. Clearly, additional studies will be required in future work to determine the mechanisms underlying this differential accumulation, but plausible consequences could be enhanced host immune-mediated clearance of the aox2 mutant at later time points and impaired virulence.

In addition to the evasion of host immune defenses, the ability to switch from yeast to filamentous morphologies is a key virulence trait of C. albicans. Periodic acid–Schiff (PAS) staining and α-Candida antibody IHC revealed numerous filamentous forms in aox2-infected kidneys (Supplementary Fig. 1b), indicating that the mutant retains the ability to undergo morphogenesis. This result is consistent with observations that AOX2 is dispensable for filamentation under in vitro tissue culture conditions that mimic the host environment (Supplementary Fig. 1c).

In addition to its role as a fungal pathogen, C. albicans is primarily a normal component of the human gut microbiota. To determine whether AOX2 is also required for commensal fitness, we tested the fitness of aox2 in a mouse model of gut colonization²². aox2-deletion and complemented strains were introduced as a 1:1 mixed inoculum directly into the stomachs of the animal model, and the relative abundance of each strain in feces was monitored every five days for the next three weeks. As shown in Supplementary Fig. 1d, the aox2 mutant maintained equivalent colonization to the complemented strain throughout the observed time-course, suggesting that AOX2 is dispensable for fitness in the commensal niche.

Aox functions in cooperation with the ETC to maintain alternative carbon source utilization

Previous studies have reported that AOX2 expression is upregulated upon phagocytosis by murine macrophages²³ and during progressive infection of kidneys in systemically infected mice²⁴. Linking this evidence with our observations of a substantial virulence defect for aox2 in a mouse model of systemic infection, we hypothesized that under host-relevant conditions, which compromise the classical ETC, C. albicans may increase Aox activity to maintain the respiratory metabolism that allows alternative carbon source utilization. To test this idea, we deleted both alleles of nuclear genes encoding subunits of each classical respiratory complex in wild-type and aox1/aox2 double homozygous deletion (aox) backgrounds. Growth was compared on solid medium containing glucose or galactose, a poorly-fermentable sugar, as the primary carbon source. As a result of compromised oxidative phosphorylation, each deletion impaired growth on YP-galactose (YPGal) medium, with the most severe defect observed in the atp2-deletion strain. Combined with the deletion of AOX genes, however, growth of the cytochrome pathway mutants (rip1, cyc1, or cox5) was completely abolished on YPGal agar. These results suggest that, in strains that lack the cytochrome pathway, Aox activity supports sufficient respiration to utilize galactose. In contrast, the growth of strains lacking NDH51 or SDH12, which retain an intact cytochrome pathway, was not further exacerbated on YPGal in the absence of Aox activity.

When cytochrome pathway mutants were grown in YPGal medium, AOX2 levels were induced to several hundred-fold higher levels than that observed in the wild-type strain (Fig. 2b). Interestingly, ndh51- and atp2-deletion strains over-expressed AOX2 in YPGal, to an even greater degree (Fig. 2b). Thus, AOX2 induction seems to be a general response to compromise of oxidative phosphorylation, rather than a specific response to loss of cytochrome pathway activity. With glucose present as a carbon source (YPD), fermentative metabolism reduces dependence on respiration. Thus, in YPD, no ETC mutants required Aox activity for growth or induced AOX2 to the same extent seen as observed in galactose medium (Fig. 2a, b).

**Fig. 2: Induction of *AOX2* supports alternative carbon source utilization upon genetic or chemical compromise of classical respiration.**

Chemical inhibition of the cytochrome pathway induces AOX2 expression

By enabling temporal control, highly specific chemical inhibitors can complement genetic techniques in revealing how ETC function contributes to fungal metabolism in different environments. Antimycin A (AA), a classical Complex III inhibitor, reduced C. albicans growth in synthetic complete medium (SC) containing poorly fermentable (galactose) or non-fermentable (glycerol, lactate) carbon sources. The growth inhibition was strongly exacerbated by the deletion of both AOX genes (Fig. 2c; Supplementary Fig. 2a). Consistent with a previous report, we found that AOX2 plays a dominant role in supporting alternative respiration¹⁵. Deletion of AOX2 resulted in the same degree of hypersensitivity to AA as deletion of both paralogs, while deletion of AOX1 alone did not alter sensitivity. Confirming the specificity of AA, dose-response assays using a structurally distinct, fungal-selective inhibitor of Complex III (Inz-5) yielded similar results in that deletion of AOX2 increased sensitivity to the compound in poorly- or non-fermentable carbon sources (Fig. 2c; Supplementary Fig. 2a)^25,26.

Use of AA allowed us to measure the kinetics of AOX2 transactivation. In YPGal, AOX2 transcripts increased sharply within 15 minutes upon addition of AA, while levels of AOX1 remained constant (Fig. 2d). A decline in AOX2 transcript level began after ~30 minutes post induction. AOX2 was induced with different kinetics in cells utilizing glycerol (YPGly): expression increased gradually to a plateau with no decline seen over a 90-minute observation period (Fig. 2d). Compared with the response seen in YPGal or YPGly, induction in YPD was transient and low in magnitude (Supplementary Fig. 2b). Lack of strong, persistent induction in YPD likely reflects a reduced dependence on respiration under fermentation-enabling conditions. To test this hypothesis, we deleted TYE7 and GAL4, which encode master transcriptional regulators of glycolytic gene expression in C. albicans²⁷. This tye7/gal4-deletion strain was more dependent on respiration for energy production than its parental wild-type strain and was hypersensitive to inhibition of respiration in YPD (Supplementary Fig. 2c). Supporting our hypothesis, AOX2 was strongly induced by AA in the tye7/gal4-deletion strain, even when utilizing glucose as a primary carbon source (Supplementary Fig. 2b). We conclude that AOX2 inducibility by AA is tightly linked to metabolic state, which in turn is contingent on environmental carbon source.

AOX1 transcript levels remained constant under all conditions tested. To determine how differential activation of the two AOX promoters dictates the abundance of their respective gene products, we compared Aox1 and Aox2 protein levels after growth in AA-containing YPGly medium. Relative Aox2 level was ≥8-fold higher than that of Aox1, when tagged and detected in the same manner (Fig. 2e). Using a strain that co-expresses RFP-tagged Aox2 and a mitochondrial marker protein, Tom70-GFP, we observed that inducible Aox2-RFP localized exclusively within mitochondria (Fig. 2f).

To determine whether an ‘inducible’ AOX1 could functionally mimic AOX2, we used the AOX2 promoter to drive AOX1 expression. The resultant hybrid gene was transactivated by AA treatment (Supplementary Fig. 2d), and fully rescued hypersensitivity to AA seen in the aox-deletion strain, akin to AOX2 driven by its native promoter (Fig. 2g).

Induction of AOX2 by ETC inhibition requires transcription factors Rtg1 and Rtg3

In filamentous fungi, transactivation of AOX gene expression is mediated by a pair of zinc cluster transcription factors (TFs)^28,29,30. In C. albicans, however, the TFs controlling AOX2 induction have not been identified. We screened a C. albicans TF-deletion library to identify mutants defective in supporting Aox-dependent growth in SC-galactose medium containing AA. Amongst the 165 TFs represented in the library³¹, rtg1- and rtg3-deletion strains showed the greatest growth defect (Fig. 3a). Deletion of only two other genes (CWT1 and DAL81) reduced growth to <20% of wild-type growth. Except for dal81-deletion, no mutants of interest had a growth defect in the absence of AA (Supplementary Fig. 3a).

**Fig. 3: Rtg1 and Rtg3 are required for *AOX2* transactivation induced by AA.**

Rtg1 and Rtg3 belong to the basic helix-loop-helix (bHLH) family of TFs. Further characterization revealed that rtg1- and rtg3-deletion mutants are hypersensitive to cytochrome pathway inhibition when utilizing diverse non-fermentable carbon sources (Fig. 3b; Supplementary Fig. 3b). Under all conditions tested, homozygous deletion of either RTG1 or RTG3 impaired growth to the same extent as homozygous deletion of both genes, consistent with their protein products functioning together as a complex^32,33. In both YPGal and YPGly, deletion of RTG3 greatly diminished the activation of AOX2 expression by AA (Fig. 3c). As a result, Aox2 protein did not accumulate in the rtg3-deletion mutant, in marked contrast to the accumulation seen in the wild-type strain (Supplementary Fig. 3c). Unlike Aox2, the level of Aox1 was not controlled by RTG3 (Supplementary Fig. 3c). Deleting RTG3 in a rip1-deletion background mirrored the effect of deleting AOX genes in that it abolished growth on YPGal agar (Supplementary Fig. 3d). Of note, AA treatment modestly increased AOX2 expression in the rtg3-deletion strain (Fig. 3c), suggesting that an Rtg1/Rtg3-independent mechanism may contribute to AOX2 induction upon inhibition of the cytochrome pathway.

In C. albicans, the Rtg1/Rtg3 complex has been implicated in regulating the expression of genes involved in multiple metabolic and signaling pathways^32,33. To confirm that defective AOX2 induction, rather than dysregulation of other Rtg1/3 target genes, renders the rtg3-deletion strain hypersensitive to AA, we replaced the native promoter of both copies of AOX2 with a tetracycline-repressible promoter (tetO) in both wild-type and rtg3-deletion backgrounds. In the absence of doxycycline (a tetracycline analog), AOX2 expression from the tetO promoter was actually greater than AA-induced expression from the native promoter (Supplementary Fig. 3e). The rtg3 mutant strain overexpressing AOX2 showed wild-type-like sensitivity to AA in SC-galactose, but hypersensitivity was restored by addition of doxycycline (and therefore repression of AOX2, Fig. 3d). Combined deletion of AOX2 and RTG3 did not have additive effects on sensitivity to cytochrome pathway inhibition by Inz-5 (Fig. 3e). We conclude that the Rtg1/Rtg3 complex supports AA-resistant alternative carbon source utilization by activating AOX2 expression.

Binding of Rtg1/Rtg3 to the AOX2 promoter is essential for AA-induced transactivation

To determine whether Rtg1/Rtg3 activates the AOX2 promoter by direct binding, we began by asking which portion(s) of the AOX2 promoter are required for activation by AA. AOX2 promoter fragments of variable length were fused to the CDR2 core promoter sequence followed by a LacZ reporter (Fig. 4a). In a reporter strain carrying the full-length AOX2 promoter construct (‘a’), LacZ expression, measured by β-galactosidase activity, was strongly induced by AA treatment in YPGly. Similar or greater induction was observed for all constructs encoding the intact −1270 to −1077 region (pAOX2 ^{−1270 to −1077}, highlighted red in Fig. 4a). Shortening this region from either end decreased inducibility. Indeed, AOX2 did not rescue AA-hypersensitivity when under control of a mutant promoter that lacks the pAOX2 ^{−1270 to −1077} fragment (Supplementary Fig. 4a). To test whether the Rtg1/Rtg3 complex can bind to this fragment in vitro, we purified recombinant complex from Escherichia coli cells that co-express Rtg1 and tagged Rtg3. The two proteins co-purified through all chromatography steps and co-eluted at a stoichiometry of ~1:1, suggesting equimolar presence of the two components in a stable complex (Fig. 4b). This complex bound double-stranded DNA encoding the pAOX2 ^{−1270 to −1077} sequence, decreasing its mobility in gel-shift assays (Fig. 4c).

**Fig. 4: An Rtg1/Rtg3 complex binds the *AOX2* promoter through multiple GTCA motifs to drive induction by AA.**

A genome-wide chromatin immunoprecipitation (ChIP) study previously reported that C. albicans Rtg1 and Rtg3 share the same consensus binding sequence³⁴. The motif at the 5′ end of this consensus sequence is highly similar to the ‘GTCAC’ motif (R box) reported to be bound by Saccharomyces cerevisiae Rtg1/Rtg3³⁵. Within the C. albicans pAOX2 ^{−1270 to −1077} fragment, we identified three GTCA motifs, which differentially contributed to Rtg1/Rtg3 binding in vitro. Interaction was strongly destabilized by mutating the GTCA motif closest to the 3′ end of the pAOX2 ^{−1270 to −1077} probe (mut.3; Fig. 4c). Disruption of the middle GTCA motif (mut. 2) also decreased binding, but to a lesser extent. A greater decrease in affinity was observed when both sites were mutated (mut. 2&3). Mutating the motif closest to the 5′ end of the probe (mut.1) did not decrease Rtg1/Rtg3 binding. When these GTCA mutations were introduced into the native AOX2 promoter in cells, the inducibility of each mutant correlated well with its relative in vitro affinity for Rtg1/Rtg3 in gel-shift assays (Fig. 4c, d). AOX2 induction driven by a promoter with all three GTCA motifs disrupted was only 10–15% of the induction seen with a wild-type promoter (Fig. 4d) and this level of induction was insufficient to support growth in SC-galactose medium containing AA (Fig. 4e).

We also directly compared the effects on AOX2 induction by AA of deleting RTG3 versus mutating the GTCA motifs. Loss of RTG3 resulted in a greater decrease in induction, suggesting that the AOX2 promoter may have additional Rtg1/Rtg3 binding site(s), which play a minor role in mediating transactivation (Supplementary Fig. 4b). Interestingly, mutating the GTCA motifs further decreased AOX2 induction in the rtg3-deletion mutant (Supplementary Fig. 4b), suggesting these motifs may also be involved in transactivation through an Rtg1/Rtg3-independent mechanism. Going forward, we refer to the −1270 to −1077 region of the AOX2 promoter as CARbox (C. albicans Antimycin Responsive box).

Inhibiting respiration induces Rtg1/Rtg3 binding to the CARbox

To learn whether increased promoter occupancy by Rtg1/Rtg3 is involved in AOX2 transactivation, we performed chromatin immunoprecipitation (ChIP) assays (Fig. 4f). The epitope tags we introduced to enable this approach did not interfere with Rtg1/Rtg3 function in activating AOX2 expression (Supplementary Fig. 4c). In cells grown in YPGal, ChIP signals were close to background levels, which were determined in a strain expressing untagged proteins (Fig. 4f). Upon exposure to AA, however, ChIP signals for both Rtg1-3XHA and Rtg3-HF increased across the AOX2 promoter concordantly. The greatest increase was at the CARbox region for both proteins (Fig. 4f). Based on in vitro and functional analyses, we predicted that Rtg1 and Rtg3 would bind as a complex to GTCA motifs within the CARbox. Indeed, mutating GTCA motifs or deleting rtg1 abolished AA-induced binding of Rtg3-HF to the CARbox (Fig. 4g, h). Substituting glucose for galactose reduced Rtg1 binding at the CARbox upon AA treatment, indicating glucose represses AOX2 induction upstream of Rtg1/Rtg3 binding to its promoter (Supplementary Fig. 4d).

In the model yeast S. cerevisiae, Rtg1/Rtg3 orthologues localize to the cytosol under basal conditions and must translocate into the nucleus to activate target gene expression³⁶. In the fungal pathogen C. albicans, however, we found different biology. Using a strain that co-expresses from their native promoters GFP-tagged Rtg3 and RFP-tagged Histone H4 (nuclear protein encoded by HHF1), we found that Rtg3-GFP co-localized with H4-RFP in the nucleus, even under non-inducing conditions (Fig. 4i). We confirmed that GFP-tagging did not interfere with Rtg3 function in regulating AOX2 expression (Supplementary Fig. 4c). We repeated these experiments using GFP-tagged Rtg1 and obtained the same results that growth in other carbon sources or AA treatment did not alter Rtg1/Rtg3 localization (Supplementary Fig. 4e, f). Additional experiments revealed that nuclear localization requires Rtg1 and Rtg3 to be in complex. Under all conditions tested, Rtg1-GFP was excluded from the nucleus in the absence of Rtg3, whereas deletion of RTG1 decreased but did not eliminate nuclear localization of Rtg3-GFP (Fig. 4i; Supplementary Fig. 4f).

Maximal transactivation requires a second TF complex consisting of Cwt1 and Zcf11

In our TF-mutant screen, two independent mutants with deletion of CWT1 were also identified as hypersensitive to AA, though to a lesser extent than rtg1- or rtg3-deletion mutants (Fig. 3a). In multiple filamentous fungi, a Cwt1 homolog acts in complex with another zinc cluster TF to regulate alternative oxidase expression^28,29,30. In C. albicans, ZCF11 encodes a predicted Cwt1 binding partner. A zcf11 mutant was not included in the TF-deletion collection that we screened. When utilizing galactose or glycerol as carbon source, we observed that a cwt1-deletion strain and a zcf11-deletion strain that we constructed showed similar degrees of hypersensitivity to Complex III inhibitors AA or Inz-5 (Fig. 5a; Supplementary Fig. 5a), which correlated with ~80% lower AOX2 transcript level after AA treatment in YPGal or in YPGly (Fig. 5b; Supplementary Fig. 5b). Double deletion of CWT1/ZCF11 phenocopied individual homozygous deletion mutants, suggesting that C. albicans Cwt1 and Zcf11 likely function as a heterocomplex to regulate Aox transactivation. Strains with either CWT1 or ZCF11 deletion were also hypersensitive to AA on agar medium containing non-fermentable carbon sources (Supplementary Fig. 5a). Of note, the cwt1-deletion mutant was defective in utilizing these carbon sources even in the absence of Complex III inhibition. The zcf11-deletion strain, however, did not show a similar phenotype, suggesting that Cwt1 has Zcf11-independent function(s) in supporting alternative carbon source utilization. Deletion of CWT1 or ZCF11 in the rtg3-deletion background virtually eliminated AOX2 induction by AA (Fig. 5b), but its effect on sensitivity to AA was masked by the overwhelming hypersensitivity caused by rtg3-deletion alone (Fig. 5a).

**Fig. 5: Cwt1 and Zcf11 comprise a transcription factor complex required for AA-induced *AOX2* transactivation.**

To further investigate how Cwt1/Zcf11 responds to AA in transactivating AOX2, we constructed strains expressing various tagged versions of these proteins from both copies of their native promoters. Modified strains showed wild-type-like growth sensitivity to AA and expressed AOX2 at levels comparable to those of untagged strains under both non-inducing and AA-inducing conditions (Supplementary Fig. 5c, d). By co-immunoprecipitation, Cwt1 was found to interact with Zcf11 not only after AA treatment, but also under non-inducing conditions (Fig. 5c). Microscopy of GFP-tagged strains revealed constitutive nuclear localization of Cwt1-GFP and Zcf11-GFP under non-inducing growth conditions (Fig. 5d). Therefore, respiration inhibition does not activate the Cwt1/Zcf11 complex by inducing heterodimerization or nuclear translocation.

We next used ChIP assays to determine whether AA treatment induces Cwt1/Zcf11 binding to the target gene-promoter. Under non-inducing conditions, Cwt1 and Zcf11 occupancy at the AOX2 promoter was similar to background but greatly increased following AA treatment (Fig. 5e). With peak occupancy at the CARbox, the ChIP binding profile of Cwt1/Zcf11 across the AOX2 promoter was similar to that of Rtg1. Deletion of ZCF11 abolished Cwt1 binding at the CARbox without changing the protein level or nuclear localization of Cwt1 (Fig. 5d, f; Supplementary Fig. 5e). Both microscopy and immunoblotting indicated that tagged Zcf11 proteins were less abundant than Cwt1 when fused to the same tags, and they became barely detectable in a cwt1-deletion background (Fig. 5d; Supplementary Fig. 5e). Collectively, these results suggest co-dependence for Cwt1 and Zcf11 binding to the CARbox to activate AOX2 transactivation. By immunoblotting, Cwt1-3XHA migrated as two major bands of the expected molecular size. The relative abundance of the two forms changed with carbon source and AA treatment shifted almost all of the Cwt1-3XHA signal to the slower-migrating form (Fig. 5g). Glucose as the carbon source or deletion of zcf11 repressed these shifts in mobility, suggesting a correlation between gel mobility and activation state for Cwt1. AA treatment also induced a mobility shift for Zcf11-3XHA, which was more subtle and independent of the carbon source (Fig. 5g). Defining the molecular basis for these changes in mobility will require additional investigation.

Cwt1/Zcf1 and Rtg1/Rtg3 bind the CARbox in a cooperative manner

Homo- or heterodimerized zinc cluster TFs, such as Cwt1/Zcf11, usually recognize a pair of CGG triplets in the same or inverted orientation spaced by nucleotides of variable length and sequence³⁷. In filamentous fungi, Cwt1/Zcf11 homologs interact with the AOX promoter through a CGG(N)₇CGG motif^28,29,38, which is absent from the CARbox or other regions of the C. albicans AOX2 promoter. Instead, the CARbox encodes three CGG triplets separated by 31 or 22 bp. Individual mutation of either of the two CGG motifs closest to the 3′ end of CARbox decreased AA-induced AOX2 expression by ~50% (Supplementary Fig. 5f). When both sites were mutated, AA-induced AOX2 expression dropped to ~15% of that driven by a wild-type promoter (Fig. 5h). By ChIP, we confirmed that AOX2 transactivation defects caused by CGG mutations correlated with reduced Cwt1 binding at the CARbox (Fig. 5h). Therefore, CGG motifs 2 and 3 are likely the binding sites for AA-activated Cwt1/Zcf11. Interestingly, each of the two CGG motifs forms a CGG(N)7CGG-like motif with the flanking sequence (underlined in Fig. 5h).

With both heterocomplexes required for maximal AOX2 transactivation, we wondered how Rtg1/Rtg3 and Cwt1/Zcf11 might interact with each other. Comparing the expression and localization of each complex before and after the deletion of the other, we found no interdependency between the two complexes for maintenance of the other’s protein level or nuclear localization (Supplementary Fig. 5g–j). Immunoprecipitation did not support inter-complex interaction to form a super-complex under either non-inducing or inducing conditions (Supplementary Fig. 5k). By ChIP, however, ~50% of Rtg1 binding was sensitive to deletion of CWT1 or ZCF11, while the other half was resistant (Fig. 5i). Similarly, binding of Cwt1/Zcf11 at the CARbox showed approximately equal contributions from Rtg3-dependent and Rtg3-independent components (Fig. 5j). Such interdependence supports a model in which the two complexes are independently activated, but bind the AOX2 promoter in a cooperative manner to achieve maximal transactivation. Consistent with this model, Cwt1-3HA in an AA-treated rtg3-deletion strain still shifted in mobility as a potential marker of activation (Supplementary Fig. 5j). Lastly, we asked how mutation of the GTCA motifs, which support Rtg1/Rtg3 binding, affect Cwt1/Zcf11 binding at the CARbox. Mutation of GTCA motif 2 partially disrupts the overlapping CGG motif 3 (Fig. 5h). Therefore, we predicted that as a consequence of losing Rtg1/Rtg3 binding and compromising a direct binding site for Cwt1/Zcf11, mutation of all three GTCA motifs would abolish Cwt1/Zcf11 binding. This prediction was confirmed by ChIP (Fig. 5k) and explains the observation that GTCA motif mutations decrease the RTG3-independent transactivation of AOX2 induced by AA (Supplementary Fig. 4b).

Cyanide treatment activates AOX2 expression through multiple mechanisms

We next examined whether other conditions that block classical respiration induce AOX2 through the same transcriptional mechanism. Similar to AA, myxothiazol, another Complex III inhibitor, requires Rtg1/3 function to potently induce AOX2 expression (Supplementary Fig. 6a). Decreasing oxygen availability by shifting cells grown in respiratory medium to an anaerobic chamber also resulted in AOX2 transactivation in a largely Rtg1/Rtg3-dependent manner (Supplementary Fig. 6b). Cyanide inhibits Complex IV and is a classical activator of AOX2 expression¹⁵. Compared with AA or the complex I inhibitor rotenone, KCN triggered the most robust AOX2 transactivation (Fig. 6a). Surprisingly, unlike rotenone or AA, KCN activated AOX2 expression primarily through an RTG3-independent mechanism. Deletion of CWT1 in an rtg3-deletion background further decreased induction by AA as expected but did not alter induction by KCN (Supplementary Fig. 6c). To confirm activation of the Rtg1/Rtg3-Cwt1/Zcf11 pathway by KCN, we assessed Rtg1 and Cwt1 binding at the AOX2 promoter by ChIP. Both transcription factors showed strong binding at the CARbox after KCN treatment (Fig. 6b), which was accompanied by a mobility shift of Cwt1 as detected by immunoblotting (Supplementary Fig. 6d). Therefore, cyanide appears to increase AOX2 expression through an additional, respiration-independent mechanism(s). Supporting this possibility, deletion of the Complex IV subunit gene COX5 did not prevent AOX2 induction by KCN (Fig. 6c). In contrast, deletion of the Complex III subunit gene RIP1 rendered AOX2 expression unresponsive to AA.

**Fig. 6: Zcf2 is the primary activator of *AOX2* in response to cyanide.**

Zcf2 acts as a cyanide-responsive regulator of AOX2 expression

Seeking an as-yet unidentified KCN-responsive regulator of AOX2, we prioritized three candidates from the zinc cluster TF family: Cta4, Stb5, and Zcf2. The first two proteins are known to increase AOX2 expression when fused with a potent transcriptional activation domain³⁹, and Zcf2 was chosen for its proposed role in supporting optimal AOX2 induction by reactive-sulfur species (RSS)⁴⁰. Measuring AOX2 induction by KCN in deletion mutants of each gene, we found that induction in YPD was not affected by deletion of CTA4 or STB5 (Supplementary Fig. 6e), but was greatly decreased in the zcf2-deletion strain (Fig. 6d). In YPGal, Zcf2 was responsible for ~85% of KCN-induced AOX2 expression (Fig. 6d), but dispensable for AOX2 induction by AA (Supplementary Fig. 6f). Therefore, Zcf2 is likely the previously unknown AOX2 transactivator that senses KCN, independent of its inhibition of Complex IV, and although less important, the Rtg1/Rtg3 pathway contributes to maximal AOX2 induction by KCN in cells utilizing galactose (Fig. 6a). Deleting RTG3 in the zcf2-deletion background further compromised AOX2 induction by KCN in YPGal (Fig. 6d). Given a report that AOX2 is required for KCN-resistant growth in medium containing alternative carbon sources¹⁵, we tested whether deleting ZCF2 and/or RTG3 to impair AOX2 transactivation would sensitize cells to growth inhibition by cyanide. Deletion of AOX2 delayed exponential growth in KCN-containing YPGly medium (Fig. 6e). The rtg3-deletion strain showed wild-type-like growth in the same medium, consistent with Rtg3’s minor contribution to AOX2 induction by KCN. Much greater KCN sensitivity was observed when ZCF2 was deleted in either a wild-type or rtg3-deletion background, suggesting that in addition to AOX2, other KCN-responsive genes are regulated by Zcf2 (Fig. 6e).

We next constructed a strain expressing tagged Zcf2 from both copies of the native promoter. It showed a ~two-fold increase in AOX2 expression under both non-inducing and cyanide-inducing conditions, suggesting that C-terminal tagging may slightly potentiate Zcf2 transcriptional activating activity, but does not impair its general function (Supplementary Fig. 6g). Tagged Zcf2 was detected at the expected molecular weight with a mobility shift seen after KCN but not AA treatment, suggesting a relationship between gel mobility and activation state (Fig. 6f). Using a strain that expresses differentially tagged Zcf2 and Rtg3, we found that under non-inducing conditions, binding of Zcf2 and Rtg3 at the AOX2 promoter were close to background level as monitored by ChIP (Fig. 6g). Both KCN and AA treatment increased Rtg3-HF ChIP signal as expected, while Zcf2 binding to the AOX2 promoter was selectively induced by cyanide. Deletion of RTG3 did not change either the level or profile of Zcf2 binding, consistent with the conclusion that Rtg3 plays a minor role in AOX2 induction by KCN (Supplementary Fig. 6h). The binding profile of Zcf2 was similar to that of Rtg3, except that the Zcf2-binding peak appeared to be broader, extending from the CARbox into the 3’ flanking region (Fig. 6g). When tested in our LacZ reporter system (Fig. 4a), the ~500 bp region that showed significant Zcf2 binding (region −1360 to −810, which is approximately covered by primer pairs 1b to 3b in Supplementary Fig. 6h) was necessary and sufficient to drive reporter transactivation in response to cyanide (Fig. 6h).

Attempts to identify the Zcf2 binding element(s) more precisely were not successful. Prior to this study, a bioinformatic analysis of the Zcf2 regulon under RSS conditions was also unable to achieve a definitive binding consensus⁴⁰. The only experimentally-confirmed element essential for Zcf2-mediated transactivation is a CCTCGG motif at a known RSS-responsive promoter⁴⁰. The −1360 to −810 region contains such a motif. However, mutating it alone or in combination with all ten of the other CGG triplets within the −1360 to −810 region did not disrupt AOX2 induction by cyanide (Supplementary Fig. 6i). Immunoprecipitation did not detect a stable interaction between 3XHA-tagged Zcf2 and 6XHis-3XFlag-tagged Zcf2 or Cwt1 under either normal or KCN-inducing conditions, suggesting that Zcf2 likely functions as a monomer (Supplementary Fig. 6j).

CARbox-region elements mediate AOX2 induction by diverse stressors to enable virulence

In animal hosts, C. albicans encounters complex nitrosative and oxidative stresses⁴¹. These types of insults, when mimicked in vitro, activate AOX2 expression^15,42,43. We found that treatment with the nitric oxide donors sodium nitroprusside (SNP) or DPTA NONOate (DTPA/NO) induced upregulation of AOX2 to a similar extent as AA, but less than KCN (Fig. 7a). Sulfite as a source of RSS also induced AOX2 expression to a level comparable to AA. Superoxide anion donor menadione only increased AOX2 expression to a level ~2% of that observed with AA. Deletion of AOX2 did not confer hypersensitivity to menadione or hydrogen peroxide (Supplementary Fig. 7). In an aox2-deletion strain that carries a re-introduced wild-type AOX2(A) allele, native-like inducibility by each stress was observed, although relative magnitudes were lower, likely because the strain only carries a single allele of AOX2 (Fig. 7a). Having characterized responses mediated by the wild-type AOX2 promoter to host-relevant chemical stresses, we compared these responses to those driven by a mutant promoter in which pAOX2 ^{−1278 to −1000} had been deleted. This promoter mutation crippled induction in response to all stressors (Fig. 7b). Therefore, pAOX2 ^{−1278 to −1000} is required not only for induction by AA/KCN, but also for induction by other challenges. Indeed, combined homozygous deletion of all three TF genes found to interact with the promoter in this region (RTG3, CWT1, and ZCF2) also impaired transactivation by all six stressors (Fig. 7b).

**Fig. 7: Defined elements within the *AOX2* promoter are required for response to specific stressors and support *C. albicans* virulence.**

Examining single deletion mutants, we found that, akin to induction by AA/KCN, maximal response to SNP, DPTA/NO, or menadione was also mediated by multiple TFs (Fig. 7b). Interestingly, DPTA/NO and SNP showed differential TF requirements for activating AOX2 expression. Induction by DPTA/NO depended on Rtg1/Rtg3 and Cwt1/Zcf11, while induction by SNP was more sensitive to the deletion of ZCF2, suggesting that SNP transactivates AOX2 through both NO- and cyanide-mediated stress. Both the Rtg1/Rtg3 and Zcf2 were essential for induction by menadione, but Cwt1/Zcf11 appeared to be dispensable. Regarding reactive-sulfur stress, single deletion of each transcription factor gene modestly decreased AOX2 induction by sulfite, but deletion of all three genes or the pAOX2 ^{−1278 to −1000} region largely abolished AOX2 induction by sulfite. All considered, we conclude that a regulatory hub consisting of the pAOX2 ^{−1278 to −1000} promoter sequence and the TFs Rtg1/Rtg3, Cwt1/Zcf11, and Zcf2 integrate signals from diverse environmental stresses to activate AOX2 expression.

In addition to the AOX2 transactivators identified here, the copper-sensing transcription factor Mac1 has previously been reported to drive AOX2 induction upon copper starvation⁴⁴. The AOX2 promoter contains multiple copies of conserved Mac1-binding elements (copper-responsive elements; CuREs)^45,46, which are located ~1 kb downstream of the CARbox. We found that deletion of these CuREs abolished AOX2 induction by copper starvation without affecting induction by AA, indicating that Mac1 mediates a distinct AOX2-inducing response (Fig. 7c).

Finally, to determine how disrupting each of the stress-selective AOX2 transactivation mechanisms identified in vitro would alter the virulence of C. albicans upon systemic infection of mice, we assembled a panel of strains created in an aox2 homozygous deletion mutant that was complemented with AOX2 under control of (1) wild-type promoter, (2) promoter with mutated GTCA motifs to selectively compromise induction through Rtg1/Rtg3 and Cwt1/Zcf11, (3) promoter with deletion of nucleotides −1278 to −1000 (pAOX2 ^{−1278 to −1000}) to disrupt Rtg1/Rtg3- Cwt1/Zcf11- and Zcf2-mediated activation, and (4) promoter with mutated CuREs to eliminate Mac1-mediated activation. Each mutant strain showed the expected condition-selective decrease in AOX2 induction compared to the wild-type-complemented strain when treated with AA, KCN, or upon growth in low-copper medium (Fig. 7d). Following intravenous infection of mice, we found that virulence of the aox2-deletion strain was restored to near-parental level by re-introduction of a single copy of AOX2 driven by its wild-type promoter or a mutant promoter with disrupted CuREs (Fig. 7e). In contrast, complementation with mutated GTCA motifs or deleted pAOX2 ^{−1278 to −1000} did not restore virulence, with a stronger defect observed for the pAOX2 ^{−1278 to −1000} deletion mutant. Overall, findings support a model in which both Rtg1/Rtg3- Cwt1/Zcf11- and Zcf2-dependent pathways contribute to C. albicans virulence by enabling AOX2 induction in response to the environmental challenges, which must be overcome to cause lethal infection (Fig. 8).

**Fig. 8: Model for transcriptional control of *AOX2* in *C. albicans*.**

Discussion

Metabolic flexibility enables microbial pathogens to invade hostile host environments and cause disease. Here, we find that the inducible expression of the alternative oxidase, AOX2 contributes greatly to the flexibility and virulence of C. albicans. Our study indicates that the virulence defect caused by loss of AOX2 expression is associated with increased leukocyte accumulation at sites of infection, either due to increased recruitment or greater survival of the responding immune cells. The difference in host response early in the course of infection when fungal burdens are comparable presumably leads to enhanced clearance of the mutant as time progresses and greater overall survival. Although the precise mechanism(s) remains to be defined, increased recognition of C. albicans by the host would be consistent with previous reports that Aox activity plays an important role in the maintenance of cell wall architecture¹⁶ and glucan masking under hypoxic conditions¹⁷.

Aox activity is found in a wide variety of organisms ranging from plants to protozoans to fungi, but not in humans⁴⁷. Although relatively inefficient in generating energy, alternative respiration allows for the direct transfer of electrons from ubiquinol to oxygen. Thus, fungi with AOX genes can bypass blockades in the classical cytochrome pathway to maintain redox balance and TCA cycle function and in turn, utilize alternative carbon sources. Besides C. albicans, the other major human fungal pathogens also have an AOX encoded in their genomes. Consistent with our findings, the virulence of C. neoformans is impaired by deletion of its AOX⁴⁸. However, previous studies with Aspergillus fumigatus found no virulence defect associated with AOX deletion⁴⁹. This disparity could be due to differences in the mouse models used to assess virulence. The A. fumigatus studies were performed in immunocompromised mice, while our studies and those with C. neoformans were performed in immunocompetent mice. At least in C. albicans, we find that fungal Aox activity reduces the accumulation of innate immune cells in organs of immunocompetent animals, which are known to generate both reactive oxygen and nitrogen species that damage classical ETC pathway components. After phagocytosis by murine macrophages, AOX2 transcript level rapidly increases even before upregulation of genes associated with filamentation, a process coupled to macrophage killing and escape²³, highlighting that induction of AOX2 is part of the primary transcriptional response of C. albicans to engulfment by host phagocytes. Alternatively, the very different pathogenetic mechanisms used by these organisms may impose differential requirements for Aox activity in enabling them to cause disease. The surprisingly complex network of transcription factors that we uncovered in C. albicans provides clues as to the specific challenges encountered during invasive infection that Aox2 may help this fungus overcome. With the relevant transcriptional activators now identified, techniques such as Calling card-Seq could be used in future work to learn where and when AOX2 expression is induced during the progression of invasive infection and in other commensal or disease models⁵⁰.

In several constitutively filamentous fungi, functional homologs of the zinc cluster transcription factor complex Cwt1/Zcf11 play an important role in the upregulation of AOX expression, as we now show for C. albicans^29,38,51. Previously unknown, however, is the role of the Rtg1/Rtg3 complex that we define here. The retrograde response pathway involving this TF complex in S. cerevisiae allows signal(s) generated by mitochondria to drive altered nuclear gene expression, but it has not been previously implicated in the control of AOX expression⁵². Within the AOX2 promoter of C. albicans, binding sites for the Cwt1/Zcf11 and Rtg1/Rtg3 complexes are in close proximity and located ~1.2 kb upstream of the transcriptional start site. Interestingly, the 1.2 kb distance between the CARbox and the AOX2 coding sequence, plus the >4 kb non-protein-coding sequence between AOX2 and the most proximal upstream gene, results in a much longer than average (557 bp) intergenic region for C. albicans⁵³. In this polymorphic fungus, genes encoding key transcription factors involved in morphogenesis and epigenetic switching between cell types are typically preceded by long 5’ intergenic regions (>7 kb) where multiple regulators can be recruited through protein-DNA interactions and protein-protein condensates to assemble highly complex transcriptional regulatory networks⁵⁴. Given the unusually long intergenic region present, coupled with the complex regulation we uncovered, it seems possible that such super-enhancer-like mechanisms could be involved in coordinating the function of the TFs we have identified binding at the AOX2 promoter.

From an organismal perspective, the complexity and magnitude of the transcriptional control we uncovered for AOX2 highlights the importance of Aox activity in enabling C. albicans virulence. Our findings begin to reveal how integration of diverse respiratory-stress stimuli can be achieved at the transcriptional level to enable adaptation of this opportunistic fungus as it shifts from harmless commensal to invasive pathogen. The insights gained deepen our molecular understanding of fungal pathogenesis, open new avenues for investigation, and could have important implications for Aox2 function as a promising, but yet to be exploited, target for antifungal drug development^16,55,56.

Methods

Ethics statement

All procedures involving animals were performed under Protocol AN189431-02C, which complies with the National Institutes of Health Guidelines for the Ethical Treatment of Animals and was approved by UCSF’s Institutional Animal Care and Use Committee.

Fungal strains and culture conditions

All strains were archived in 25% glycerol at −80 °C for long-term storage and freshly streaked on YPD agar medium within 1-week prior to experiments. Strains grown on agar plates were maintained at ambient temperature without refrigeration. In this study, YP-based media contained 1% yeast extract, 2% peptone, 0.2 mM uridine, and one of the following carbon sources: 2% glucose (YPD), 2% galactose (YPGal) or 3% (v/v) glycerol (YPGly). One liter of synthetic complete medium (SC) contained 6.7 g yeast nitrogen base (YNB; with ammonium sulfate), 2 g Drop-out mix without uracil, 0.2 mmol uridine, and one of the following carbon sources: 20 g glucose, 20 g galactose, 30 mL glycerol, 20 g sodium acetate, 20 g casamino acid, 20 g sodium citrate or 30 g sodium lactate. One liter of low-copper SC medium contained 1.7 g YNB (without ammonium sulfate, ferric chloride, or copper sulfate; US Biologicals), 1 g monosodium glutamate hydrate, 2 g Drop-out mix without uracil, 0.2 mmol uridine, 20 g galactose, and 2.5 µmol ferric chloride. To induce copper starvation, strains were grown to saturation in low-copper SC medium and diluted into fresh low-copper medium supplemented with 50 µM bathocuproinedisulfonic acid (BCS) for ~5–6 h. Sulfite treatment was performed in acidified YPGal (adjusted to pH 4.0 with HCl). Cells were grown in buffered YPGal (adjusted to pH 7.2 by the addition of 25 mM HEPES-KOH) before treatment with sodium nitroprusside or DPTA NONOate. Solid media contain 1.5–2% agar.

Systemic candidiasis model in mice

Experiments were performed with 8- to 10-week-old female BALB/c mice (n = 5 or n = 8 per experimental group) from Charles River Laboratories (Strain code 028). Systemic infection with C. albicans strains was performed by injection of 1 × 10⁵ Colony-Forming Units (CFU) of mid-log-phase yeast into the retrobulbar sinus under isoflurane anesthesia. Animals were provided with fresh water and chow ad libitum, Housing was performed under a 12:12 light: dark cycle in a well-ventilated room (10-15 air changes per hour) with stabilized temperature (68–79 degrees Fahrenheit) and humidity (30-70%). Infected animals were assessed at least once a day for signs of significant morbidity (decreased motor activity, respiratory distress, or a body condition score (BCS) of 2 or less). To minimize suffering and guarantee humane euthanasia, animals that displayed any of these criteria were immediately euthanized by two independent procedures (administration of CO2, followed by cervical dislocation or bilateral thoracotomy).

Determination of fungal burden in organs

Mice (n = 5) were infected with 1 × 10⁵ CFU with the aox2-deletion mutant or the complemented strain via the retrobulbar sinus. The kidney, liver, spleen, and brain were collected on Day 3 post-infection. Organs were weighted and homogenized in 1 mL of saline solution (0.9%). Serial dilutions were performed and plated on Sabouraud agar. CFUs were counted after 2 days of growth at 30 °C and were normalized by the organ’s weight.

Histological analysis of infected kidneys

Kidneys from mice infected with aox2-deletion mutant or the complemented strain were collected on Day 3 post-infection and formalin-fixed/paraffin-embedded using standard methods. Serial sections (4 µm) taken at three non-contiguous levels (>70 µm between levels) were stained by routine H&E and PAS methods. For IHC, α-Candida (1:1000, Abcam ab53891) and α-CD45 (1:1000, Invitrogen RA3-6B2) antibodies were used followed by detection with diaminobenzidine (DAB) according to manufacturer’s recommendations (ImmPACT DAB Substrate Kit, Peroxidase-Biolynx, Vector Laboratories, SK-4105). Slides were imaged on a 3DH Pannoramic Slide Scanner at 40X magnification (bright field).

Commensalism model in mice

The mouse model of C. albicans commensalism was performed as previously described^57,58. Female BALB/c mice (n = 4, 8–10-week-olds) were singly housed and treated with penicillin 1500 un/ml and streptomycin 2 mg/mL in drinking water starting 7 days before gavage and continuously for the remainder of the experiment. Feces were plated on Sabouraud agar, and LB agar to monitor for fungal and bacterial growth. Mice were infected with 10⁸ CFUs of a 1:1 mixture of the aox2-deletion mutant and the complemented strain in 0.9% saline by gavage. C. albicans was recovered from the inoculum and host feces every 5 days for 3 weeks, and strain quantification was performed by qPCR using primers listed in Supplementary Data 2.

Strain construction

Detailed information regarding the strains used in this study, including names, genotypes, and figure panel(s) in which each strain was presented are provided in Supplementary Data 1. The transient CRISPR system developed by Min et al. ⁵⁹. was used to tag or delete both alleles of C. albicans genes of interest. The tagging system described by Zhang et al. ⁶⁰. was used to modify C. albicans to express proteins with C-terminal tags from their endogenous promoters. Additional details regarding strain and plasmid construction are available upon request.

Dose-response and spotting growth inhibition assays

Strains grown overnight in YPD were diluted to ~50,000 cells/mL in the appropriate medium. 100 µL (or 20 µL) of the diluted culture was mixed with an equal volume of medium supplemented with a given inhibitor at the specified range of concentrations in 96- or a 384-well plate format. Growth (OD₆₀₀) in SC-glucose or SC-galactose was measured after incubation at 30 °C for 48 h. Ten-fold higher concentration inocula were used when glycerol, acetate, lactate, or citrate was the carbon source and growth was measured after 72-hour incubation. Unless otherwise specified, growth (OD₆₀₀) in the presence of inhibitor was normalized to growth in a compound-free medium. All dose-response assays were performed at least twice with comparable results. The mean of the technical duplicates from one representative experiment was presented in heatmap format visualized by Java TreeView, version 1.1.6r4 (http://jtreeview.sourceforge.net). For spotting growth assays on agar medium, strains were grown to saturation in tubes under shaking conditions or statically in a 96-well plate. Ten-fold serial dilutions of cultures were made in water and spotted onto agar plates using a multi-channel pipettor or a 96-well replicator.

Compound treatments

The growth medium, compound concentration, and duration of each treatment are specified for each experiment in the Figure Legend. Strains were grown overnight in the same medium in which treatment would be performed. The saturated culture was diluted in fresh medium (1: 50 for YPGly, 1:100 for YPGal, and 1:200 for YPD) and sub-cultured for ~4–5 h before treatment. Antimycin A, rotenone, myxothiazol, and Inz-5 were added to the indicated final concentration from 10 mM DSMO stocks stored at −20 °C. KCN solution (1 M) was kept in dark at room temperature. Fresh stocks were prepared every 2-3 months. Menadione in DMSO (0.25 M) and sodium sulfite in water (0.1 M) were freshly prepared for each experiment. Sodium nitroprusside dihydrate (0.5 M in water) and DPTA NONOate (0.1 M in YPGal medium buffered to pH 7.2 by 25 mM HEPES-KOH) were prepared ~10 min before being added to fungal cultures.

Quantitative reverse transcription PCR

Total RNA preparation and reverse transcription were performed as previously described⁶¹. Reactions containing cDNA reverse-transcribed from 1 µg RNA were 1:80 diluted in water and then diluted samples (4 µL) were used to assess relative gene expression in a 10 µL SYBR green quantitative PCR reaction (Fast SYBR™ Green Master Mix) with ACT1 level as the internal control. Sequences of the qPCR primers used in this study are listed in Supplementary Data 2.

β-galactosidase activity assays

Strains carrying each LacZ reporter construct were grown to early-log-phase in YPGly medium and aliquoted into a 96-well plate (150 µL/well). After the addition of 10 µM AA (or DMSO vehicle control), cells were grown for ~4 h at 30 °C under shaking conditions. β-galactosidase activity was then measured using a Gal-Screen^TM kit by mixing an equal volume of cell culture (30 µL) and Buffer B (provided in the kit) in a 384-well plate. Luminescent signals were recorded by a Tecan plate reader (controlled by Magellan 7.2) every 15 min for 90 min during incubation at 30 °C and peak signal, which was typically reached after 60-75 min, was used to compare induction of LacZ after normalization to the OD₆₀₀ of the input culture.

Purification of recombinant Rtg1/Rtg3-6Xhis complex from E. coli

Codon-optimized sequences were inserted between the NcoI/SacI (Rtg1) and the NdeI/XhoI (Rtg3-6XHis) sites of the pColaDuet vector (Novagen) for co-expression in E. coli (Supplementary Data 3). A BL21(DE3) expression strain carrying the pColaDuet-Rtg1-Rtg3X6his plasmid (pLC1452) was grown in four liters of LB+Kan liquid medium (1% Tryptone, 0.5% yeast extract, 1% NaCl and 50 µg/mL kanamycin) to OD₆₀₀ ~ 0.6 at 37 °C. Expression of the recombinant protein was induced for ~4 h at 30 °C after addition of 1 mM IPTG. Cells were collected by centrifugation at 12,000 × g for 15 min and resuspended in 35 mL (per liter culture) lysis buffer (50 mM NaPi, pH 8.0, 300 mM NaCl and 5 mM imidazole) and sonicated for 20 min (10 sec on; 10 sec off) at 30% amplitude using a Misonix S-4000 dual horn sonicator with 3/4-inch probes. Lysates were then clarified by centrifugation at 35,000 × g for 25 min. The supernatant was filtered through 0.45 µm PVDF membrane and incubated with Ni-NTA agarose (2 mL bed volume) for 1 h. Beads were collected into a chromatography column and washed with 40 mL of wash buffer (50 mM NaPi pH 8.0, 600 mM NaCl, and 15 mM imidazole). Bound proteins were eluted twice with 2.5 mL elution buffer (50 mM NaPi pH 8.0, 300 mM NaCl, and 250 mM imidazole). After buffer-exchange on a PD-10 desalting column to T150 buffer (25 mM Tris-HCl pH 7.5, 150 mM NaCl, and 5% glycerol), the product was applied to HiTrap Heparin HP (5 mL; GE) and eluted by increasing concentrations of NaCl (0.15 to 1 M). Peak fractions were combined and re-concentrated on ~0.6 mL bed volume of Ni-NTA agarose. This sample was resolved on HiLoad 16/600 Superdex 200 pg size-exclusion column (SEC; running buffer: 25 mM HEPES-NaOH pH 7.8, 300 mM NaCl, and 5% glycerol). All centrifugation and Ni-NTA purification steps were performed at 4 °C, while heparin and size-exclusion chromatography were performed at ambient temperature. All buffers were supplemented with a protease inhibitor cocktail immediately prior to use (1 mM PMSF, 2 mM benzamidine HCl, 0.6 μM leupeptin, and 2 μM pepstatin A).

Gel-shift assays

The concentration of recombinant Rtg1/Rtg3-6XHis in peak SEC elution factions was estimated using a molar extinction coefficient of 47,220 M⁻¹ cm⁻¹ at 280 nm. Serial dilution of the recombinant proteins (Mix A) in SEC running buffer was made from an ~7.5 μM fraction. DNA probes were amplified from plasmids encoding WT or mutant AOX2 promoters using ExTaq polymerase, purified on QuickSpin columns, and stored as 20 nM stocks in EB buffer. The TCA triplet closely following the GTCA motif 1 or motif 3 was disrupted with the associated GTCA motif to generate mutant vectors and probes. Sequences of the probes and the full-length AOX2(A)/(B) promoters cloned from wild-type strain SN95 are provided in Supplementary Data 2 and 3, respectively. Mix B was made by diluting the probes to 2.67 nM in a buffer containing 25 mM HEPE-NaOH pH 7.2, 1 mM EDTA, and 5% glycerol. 5 µL of Mix A and 15 µL of Mix B were combined in a low-binding 1.5 mL tube (VWR) and incubated for ~15 min at ambient temperature. Samples were directly loaded on a 5% TBE-glycerol acrylamide gel (5% acrylamide/bis-acrylamide (29:1), 0.5× TBE and 2.5% glycerol), which had been pre-run for ~30 min at 100 V. After electrophoresis at 10 mA for ~80 min in 0.5XTBE + 2.5% glycerol at ambient temperature, gels were stained in running buffer containing 1× SYBR Green for 20 min and imaged using ChemiDoc (Bio-Rad) after three rinses in distilled water.

Immunoprecipitation

Strains, growth media, and compound treatment for IP experiments are specified in each Figure Legend. Routinely, cultures (100 mL) in YPGal before and after AA or KCN treatment were harvested by centrifugation at 3000 × g for 2 min in 50 mL conical tubes. Cell pellets were washed in 1 mL of F300 buffer (50 mM HEPES-KOH pH 7.2, 300 mM KOAc, and 10% glycerol) and stored at −80 °C. To prepare whole cell lysate, each pellet was resuspended in 1 mL F300 and disrupted by bead-beating (2 min on; 1 min rest on ice) for three cycles. Crude lysates were mixed with a half-volume of F300 supplemented with 0.06% IGPAL-CA630 and sonicated (20 sec × 4) at 30% amplitude using a 0.63-cm diameter probe. Sonicated lysates were treated with ~30 U Benzonase (Millipore) for 20 min after the addition of MgCl₂ to the final centration of 2 mM and clarified by centrifugation at 20,000 × g for 20 min. Total protein (~6 mg) was incubated with ~6 µg of anti-Flag (or anti-HA) antibody for 90 min on a revolving rotator. Protein G Dynabeads (40 µL) were added to each IP and incubated for 90 min. Bead-bound antibody-protein complexes were captured using a magnetic rack and washed with F300 + 0.02% IGEPAL-CA630 × 3 before boiling in 1× SDS-PAGE loading buffer (40 µL). Proteins were resolved on 10% precast SDS-gels and blots probed with HRP-conjugated anti-Flag antibody (A8592; Sigma; 1:3000) or anti-HA antibody (3F10; Roche; 1:3000) followed by HRP-conjugated goat anti-Rat secondary antibody (A10549 Invitrogen; 1:7000). All F300 buffers were supplemented immediately prior to use with 1× PhosSTOP (Roche) and 1× Complete inhibitor cocktails (Roche) to inhibit phosphatase and proteinase activity.

Whole-cell lysate preparation under denaturing conditions and immunoblotting

Whole cells lysates for assessing relative protein level or gel mobility were prepared using a TCA-precipitation-based method as previously described with slight modification⁶². Precipitated protein pellets were rinsed in cold acetone and resuspended in a modified loading buffer (40 mM Tris-HCl pH 6.8, 8 M urea, 5% SDS, 0.1 M EDTA), which did not contain β-mercaptoethanol or bromophenol blue. Samples were diluted 1:20 in 1% SDS to measure protein concentration by DC protein assay kit using BSA as a standard. Samples were diluted to the same protein concentration and mixed with 1/5 volume of the modified loading buffer supplemented with 0.6% (v/v) β-mercaptoethanol and ~0.3 mg/mL bromophenol blue. Total protein (~30 µg) in samples was resolved on 10% SDS-gels and blots probed using anti-Flag (F3165; Sigma; 1:3000) or anti-HA (3F10; Roche; 1:3000) primary antibody and the corresponding HRP-conjugated secondary antibody. Blots were developed using Clarity^TM ECL substrate (Bio-Rad) and chemiluminescent signals captured by ChemiDoc Imager (Bio-Rad).

Chromatin immunoprecipitation assay (ChIP)

ChIP assays were performed as previously described with minor modifications⁶². Cells were fixed in 1% formaldehyde for 20 min at room temperature with agitation. Crude lysates recovered after bead-beating were successively sonicated using a probe (0.63-cm diameter; 20 sec × 4; 30% amplitude) and then a water bath sonicator (Bio-Disrupter; High setting; 5 min × 4; 30 sec on/30 sec off). Antibodies used for anti-Flag and anti-HA ChIP were F3165 (Sigma) and sc-7392 X (Santa Cruz) respectively. The abundance of the DNA fragments of interest in input samples and IP products were assessed by qPCR using primers listed in Supplementary Data 2.

Fluorescence microscopy

Strains and growth conditions for each microscopy experiment are specified in Figure Legends. Routinely, ~ 1 mL of log-phase cells was pelleted by centrifugation. After washing in 500 µL PBS twice, cells were resuspended in ~150 µL PBS and immediately imaged under a Zeiss Axio Imager MI microscope (Carl Zeiss) with an X-cite series 120 light source for fluorescence. Images were captured and processed by the ZEN pro program (Zeiss).

Statistics and reproducibility

All RT-qPCR, ChIP-qPCR experiments, and dose-response assays were performed at least twice and/or using independent transformants to ensure reproducibility. Except where indicated in specific figure legends, data are presented as the mean (SD) of technical replicates from one representative biological experiment whenever major (>5-fold differences between test and control groups) were observed. Data for the corresponding biological replicate confirming reproducibility are provided in the source data file. When less than twofold differences between test and control groups were observed, data from three biological replicates are presented and significance was determined by unpaired, two-tailed Student’s t test. GraphPad Prism (7.04) was used to perform Kaplan–Meier survival analysis, generate graphical displays, and perform tests of significance using Log-rank (Mantel-Cox) tests. Microscopic analyses of fungal morphology (Supplementary Fig. 1c) or localization of GFP/RFP fusion proteins (Figs. 2f, 4i, 5d; Supplementary Figs. 4e, f, 5g, i) were performed twice using biologically distinct samples to ensure reproducibility. Representative fields containing multiple (≥4) cells from one experiment were presented. Results of gel-shift assays (Fig. 4c), immunoprecipitation (Fig. 5c; Supplementary Fig. 5k, 6j), and immunoblotting comparing protein abundance or mobility (Figs. 2e, 5g, 6f; Supplementary Fig. 3c, 5e, h, j, 6d) were representative of two biological replicates. For purification of the recombinant Rtg1/Rtg3-6XHis complex (Fig. 4b), multiple batches of Ni-NTA products showed a similar elution profile from the Heparin column. Size-exclusion chromatography was performed once using pooled Heparin eluates.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Source data are provided in this paper, which includes all raw data reported within this manuscript. Source data are provided in this paper.

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Acknowledgements

We thank all current and past Cowen lab members for helpful discussions. L.E.C. is supported by the Canadian Institutes of Health Research (CIHR) Foundation grant (FDN-154288), a National Institutes of Health (NIH) R01 grant (R01AI127375), and a CIFAR Catalyst Award (CP21-042); L.E.C. is a Canada Research Chair (Tier 1) in Microbial Genomics & Infectious Disease and co-Director of the CIFAR Fungal Kingdom: Threats & Opportunities program. Z.L. was supported in part by a fellowship from the University of Toronto PRiME Initiative.

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Department of Molecular Genetics, University of Toronto, Toronto, ON, Canada
Zhongle Liu, Saif Hossain, Sean D. Liston, Nicole Robbins, Luke Whitesell & Leah E. Cowen
UCSF Department of Microbiology & Immunology, San Francisco, CA, USA
Pauline Basso & Suzanne M. Noble
UCSF Department of Medicine, Division of Infectious Diseases, San Francisco, CA, USA
Suzanne M. Noble

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Zhongle Liu
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Pauline Basso
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Saif Hossain
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Sean D. Liston
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Nicole Robbins
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Luke Whitesell
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Suzanne M. Noble
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Leah E. Cowen
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Contributions

Z.L., L.W., and L.E.C. conceptualized the study. L.E.C. and S.M.N. obtained funding and provided resources for the study. Z.L., P.B., and S.D.L. conducted investigations. Z.L., P.B., and S.H. performed analysis and visualization. N.R., L.W., S.M.N., and L.E.C. supervised the study. Z.L., L.W., N.R., and L.E.C. wrote the draft manuscript, and all authors reviewed and edited the final manuscript.

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Correspondence to Suzanne M. Noble or Leah E. Cowen.

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Competing interests

L.E.C. and L.W. are co-founders and shareholders in Bright Angel Therapeutics, a platform company for the development of novel antifungal therapeutics. L.E.C. is a Science Advisor for Kapoose Creek, a company that harnesses the therapeutic potential of fungi. All other authors declare no competing interests.

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Liu, Z., Basso, P., Hossain, S. et al. Multifactor transcriptional control of alternative oxidase induction integrates diverse environmental inputs to enable fungal virulence. Nat Commun 14, 4528 (2023). https://doi.org/10.1038/s41467-023-40209-w

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Received: 11 July 2022
Accepted: 17 July 2023
Published: 27 July 2023
DOI: https://doi.org/10.1038/s41467-023-40209-w

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