The C. albicans, C. glabrata, and S. cerevisiae strains used in this study are listed in Supplementary Table 1 . All strains were stored as frozen stocks containing 20% glycerol at -80°C and sub-cultured on YPD agar plates (1% yeast extract, 2% peptone, 2% glucose, 2% agar) at 30°C for 2 days. Strains were routinely grown in YPD liquid medium at 30°C overnight with shaking at 180 rpm.

C. albicans Casky1Δ andCasky2Δ (orf19.35) deletion mutants were taken from the existent mutant library of non-essential uncharacterized predicted protein kinases (Ramírez-Zavala et al., 2022). All other C. albicans deletion mutants and complemented strains were constructed as described previously (Ramírez-Zavala et al., 2017). C. albicans PTR22 overexpression strains were generated using the ApaI-SacII fragment from plasmid pPTR22E1 to integrate PTR22 under the control of the ADH1 promoter in the wild-type strain SC5314 and the sky2Δ mutants as described previously (Reuss et al., 2004; Dunkel et al., 2013). The correct genomic integration of all constructs was confirmed by Southern hybridization with the upstream and downstream flanking sequences. All strains and primers are listed in Supplementary Table 1 .

The generation of Cgsky1Δ was conducted with a PCR-based Gibson Assembly cloning approach (NEB) according to the manufacturers’ protocol. Purified PCR fragments for the puC19 vector backbone, the CgSKY1 5’ flank (~1000 bp), a barcoded nourseothricin resistance cassette (NAT1) with constant flanking regions (U1 and D1) derived from the mutant 7G6 from Schwarzmüller et al. (2014), and the CgSKY1 3’ flank (~1000 bp) were fused into one vector. The deletion construct was verified by sequencing, then PCR-amplified and used to transform the C. glabrata wild-type strain ATCC2001 by a modified heat shock method (with 45°C heat shock for 15 min) (Sanglard et al., 1996). The transformants were plated onto YPD agar plates containing 200 µg/ml nourseothricin and positive knockout strains were verified by control PCRs. All strains and primers are listed in Supplementary Table 1 .

The S. cerevisiae wild-type strain BY4741 (Y00000) and the Scsky1Δ mutant (YMR216C) were obtained from Euroscarf (www.euroscarf.de). Both strains were transformed with the plasmid pHLUM (Addgene, Massachusetts, USA) by a LiAc/SS carrier DNA/PEG method to restore the non-auxotrophic strain background as described previously (Gietz and Schiestl, 2007).

The alignment was generated using ‘MUSCLE Alignment’ implemented in Geneious Prime (v2020.1.1.) using default settings. Sequence data originates from GenBank (NP_003128, NP_013943) and the Candida Genome Database (C1_06090C, C2_06600W, CAGL0F03905g). The data presented in Figure 1 was exported from Geneious Prime. The detailed alignment of the kinase domain is illustrated by applying the R-package ‘ggmsa’ (by Guangchuang Yu, https://CRAN.R-project.org/package=ggmsa) using the color scheme ‘Clustal’.

The C. albicans wild-type strain SC5314 and strain A of the Casky1Δ and Casky2Δ mutants ( Supplementary Table 1 ) were pre-grown on YPD plates. High-throughput phenotypic screen was performed using Phenotype MicroArrays for microbial cells (PM) plates, reagents, and devices according to the manufacturers’ instructions (Biolog, Inc., USA). Briefly, C. albicans cells were scraped from YPD agar plates and the cell number was adjusted in sterile dH₂O to 62% transmittance as measured by a turbidimeter (Biolog, Inc., USA). The cells were added to inoculating fluid IFY-0 base (1.2x), redox dye mix D (75x) (Biolog Inc., USA), and further supplemented with potassium phosphate, sodium sulfate and either glucose or glutamic acid (Sigma-Aldrich). 100 µl of the respective mixture was added to each well of PM plates for fungi (PM1–10 and PM21–25) to test for metabolic activity in the presence of different carbon sources, nitrogen sources, supplements, and chemicals. Plates were incubated at 37°C in an OmniLog multiple plate reader (Biolog, Inc., USA). Metabolic activity was determined by a colorimetric reduction of the redox dye and kinetically measured every 15 min at an optical density OD of 750 nm for 24–48 h. Each experiment was performed in duplicates.

For analysis, the respective negative control was substracted from the growth signals in each Phenotype MicroArray. Negative values were replaced by zero, if present. Next, each array’s growth signals were categorized into two groups: exponential growth (active growth) and no exponential growth (non-active growth), as proposed by Vehkala et al. (2015). In brief, a data fitted logistic curve represented exponential growth, while a line without showing exponential growth phase characteristics was interpreted as non-active growth in the investigated time frame. The method was repeated for each replicate separately. Hence, a given substrate’s growth signal was detected as exponential growth in two replicates or in no replicates if the analysis identified the same behavior for both replicates. Expoential growth in one replicate was identified if not all replicates for a given tested metabolite showed active growth. Due to low sample numbers, growth differences were assessed by comparing log₂(fold changes) of the last time point per substrate per phenotypic microarray assay. Grouping the growth signals was implemented in R version 3.6.0 with the pipeline proposed by Vehkala et al., (Vehkala et al., 2015) built upon the opm package version 1.3.77 (Vaas et al., 2013).

YPD overnight cultures of the wild type and mutant strains were centrifuged (4,000 × g, 5 min) and washed twice with dH₂O. Susceptibility to osmotic stress and stress caused by polyamines was tested by spot dilution assays on YPD agar plates as described in the respective figure legends. Briefly, strains were adjusted to an optical density of 600 nm (OD₆₀₀) of 1.0 and 5 µl of each 10-fold serial dilutions was spotted onto the respective plates and incubated for 2 days at either 30°C or 37°C.

Growth curves in liquid media were performed in YCB medium (11.7 g/l yeast carbon base, pH 5.0) containing 10 mM of the respective dipeptides. YPD overnight cultures of the respective C. albicans strains were centrifuged (4,000 × g, 5 min) and washed twice with dH₂O. Strains were adjusted to an OD₆₀₀ of 0.01 in 12 ml YCB medium containing 10 mM of the respective dipeptides and incubated for 48 h at 37°C shaking at 180 rpm. The OD₆₀₀ was measured after 24 h and 48 h. In addition, growth curves were performed in SD medium (0.17% (w/v) yeast nitrogen base without ammonium sulfate and amino acids, 0.5% (w/v) ammonium sulfate, 2% (w/v) glucose, pH 5.0) supplemented with 140 µg/ml Nikkomycin Z (Sigma-Aldrich) or 130 µg/ml Polyoxin D (Sigma-Aldrich) to test for sensitivity to these antibiotics. Strains were adjusted to an OD₆₀₀ of 0.02 in 100 µl of the respective medium in 96-well plates. Plates were incubated at 37°C in an Infinite 200 Pro plate reader (Tecan, Germany) and absorbance was measured every 20 min at an optical density of 600 nm for 24 h or 48 h. Each strain was measured in triplicates and the means and standard deviations were calculated.

YPD overnight cultures of the wild-type strain SC5314 and mutant strains were adjusted to an OD₆₀₀ of 0.1 in 900 ml YPD medium and grown for 4 h at 37°C. Cells were disrupted by using mortar and pestle with liquid nitrogen. Cell debris were homogenized in lysis buffer (1% (w/v) SDS, 150 mM NaCl, 100 mM TEAB (triethyl ammonium bicarbonate), one tablet each of cOmplete Ultra Protease Inhibitor Cocktail and PhosSTOP). After addition of 0.5 µl Benzonase nuclease (250 U/μl) the samples were incubated at 37°C in a water bath sonicator for 30 min. Proteins were separated from insolubilized debris by centrifugation (15 min, 18000 × g). Each 6 mg of total protein per sample was diluted with 100 mM TEAB to gain a final volume of 4 ml. Subsequently, cysteine thiols were reduced and carbamidomethylated in one step for 30 min at 70°C by addition of 120 µl of 500 mM TCEP (tris(2-carboxyethyl)phosphine) and 120 µl of 625 mM 2-chloroacetamide (CAA). The samples were further cleaned up by methanol-chloroform-water precipitation using the protocol of Wessel and Flügge (Wessel and Flügge, 1984). Protein precipitates were resolubilized in 5% trifluoroethanol of aqueous 100 mM TEAB and digested overnight (18 h) with a Trypsin+LysC mixture (Promega) at a protein to protease ratio of 33:1. Each sample was divided in 6 × 1 mg used for the phosphopeptide enrichment and 200 µg initial protein used as reference for proteome analysis. Samples were evaporated in a SpeedVac. The reference proteome sample was resolubilized in 50 µl of 0.05% TFA in H₂O/ACN 98/2 (v/v) filtered through Ultrafree-MC 0.2 µm PTFE membrane spin filters (Merck-Millipore). The filtrate was transferred to HPLC vials and injected into the LC-MS/MS instrument for further analysis.

Phosphopeptides were enriched by using TiO₂+ZrO₂ TopTips (Glygen Corp., Columbia, MD, USA). TopTips were loaded with 1 mg protein isolate using 6 TopTips per biological replicate after equilibration with 200 µl Load and Wash Solution 1, LWS1 (1% trifluoroacetic acid (TFA), 20% lactic acid, 25% acetonitrile (ACN), 54% H₂O). TopTips were centrifuged at 1500 rpm (~200 × g) for 5 min at room temperature. After washing with 200 µl LWS1, the TiO₂/ZrO₂ resin was washed with 25% ACN and subsequently the phosphopeptides were eluted with 200 µl NH₃· H₂O (NH₄OH), pH 12. The alkaline solution was immediately evaporated using a vacuum concentrator (Eppendorf). The phosphoproteome samples were resolubilized in 50 µl of 0.05% TFA in H₂O/ACN 98/2 (v/v) filtered through Ultrafree-MC 0.2 µm PTFE membrane spin filters (Merck-Millipore). The filtrate was then transferred to HPLC vials and injected into the LC-MS/MS instrument for further analysis.

Each sample was measured in triplicates (3 analytical replicates of 3 biological replicates of a reference proteome fraction and a phosphoproteome fraction). LC-MS/MS analysis was performed on an Ultimate 3000 nano RSLC system connected to a QExactive HF mass spectrometer (Thermo Fisher Scientific, Waltham, MA, USA). Peptide trapping for 5 min on an Acclaim Pep Map 100 column (2 cm × 75 µm, 3 µm) at 5 µl/min was followed by separation on an analytical Acclaim Pep Map RSLC nano column (50 cm × 75 µm, 2 µm). Mobile phase gradient elution of eluent A (0.1% (v/v) formic acid in water) mixed with eluent B (0.1% (v/v) formic acid in 90/10 acetonitrile/water) was performed using the following gradient for the more hydrophilic phosphoproteome samples: 0–5 min at 4% B, 15 min at 7% B, 50 min at 10% B, 100 min at 14% B, 150 min at 25% B, 190 min at 60% B, 205–215 min at 96% B, 215–240 min at 4% B. The reference proteome gradient was as follows: 0–4 min at 4% B, 10 min at 7% B, 50 min at 12% B, 100 min at 16% B, 150 min at 25% B, 175 min at 35% B, 200 min at 60% B, 210–215 min at 96% B, 215–240 min at 4% B.

Positively charged ions were generated at a spray voltage of 2.2 kV using a stainless-steel emitter attached to the Nanospray Flex Ion Source (Thermo Fisher Scientific). The quadrupole/orbitrap instrument was operated in Full MS/data dependent MS2 Top15 mode. Precursor ions were monitored at m/z 300–1500 at a resolution of 120,000 FWHM (full width at half maximum) using a maximum injection time (ITmax) of 120 ms and an AGC (automatic gain control) target of 3 × 10⁶. Precursor ions with a charge state of z=2–5 were filtered at an isolation width of m/z 1.6 amu for further HCD fragmentation at 30% normalized collision energy (NCE). MS2 ions were scanned at 15,000 FWHM (ITmax=100 ms, AGC= 2 × 10⁵) using a fixed first mass of m/z 120 amu. Dynamic exclusion of precursor ions was set to 30 s and the minimum AGC target for Precursor ions selected for HCD fragmentation was set to 1e3. The LC-MS/MS instrument was run by Chromeleon 7.2, QExactive HF Tune 2.8 and Xcalibur 4.0 software.

Tandem mass spectra were searched against the UniProt database (2021/07/19) (YYYY/MM/DD); (https://www.uniprot.org/proteomes/UP000000559) of Candida albicans SC5314 using Proteome Discoverer (PD) 2.4 (Thermo) and the algorithms of Mascot 2.4.1 (Matrix Science, UK), Sequest HT (version of PD2.4), MS Amanda 2.0, and MS Fragger 3.2. Two missed cleavages were allowed for the tryptic digestion. The precursor mass tolerance was set to 10 ppm and the fragment mass tolerance was set to 0.02 Da. Modifications were defined as dynamic Met oxidation, phosphorylation of Ser, Thr, and Tyr, protein N-term acetylation as well as static Cys carbamidomethylation. A strict false discovery rate (FDR) < 1% (peptide and protein level) and a search engine score of > 30 (Mascot), > 4 (Sequest HT), > 300 (MS Amanda) or > 8 (MS Fragger) was required for positive protein hits. The Percolator node of PD2.4 and a reverse decoy database was used for q value validation of spectral matches. Only rank 1 proteins and peptides of the top scored proteins were counted. Label-free protein quantification was based on the Minora algorithm of PD2.4 using the precursor abundance based on intensity and a signal-to-noise ratio > 5. Normalization was performed by using the total peptide amount method. Imputation of missing quan values was applied by using abundance values of 75% of the lowest abundance identified per sample. For the reference proteome analysis used for master protein abundance correction of the phosphoproteome data, phosphopeptides were excluded from quantification. Differential protein abundance was defined as a fold change of > 4, ratio-adjusted p-value < 0.05 (p-value/log₄ratio) and at least identified in 3 of 3 replicates. Differential phosphopeptide abundance was defined as a fold change of > 4, ratio-adjusted p-value < 0.05 (p-value/log₄ratio) and at least identified in 2 of 3 replicates. The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE (Perez-Riverol et al., 2019) partner repository with the dataset identifier PXD027612.

According to the Candida Genome Database (CGD), the C. albicans genome encodes two predicted SR protein kinases, based on sequence similarity to the sole SR-like protein kinase in S. cerevisiae, ScSky1. These orthologs correspond to the proteins encoded by C1_06090C (48.5% identity to ScSky1) and C2_06600W (46.5% identity to ScSky1), named in this study CaSky1 and CaSky2, respectively. To ensure that these are the only SRPKs in the pathogenic fungus C. albicans and to identify homologs in other pathogenic Candida spp., we performed a protein-protein Basic Local Alignment Search Tool (BLASTp) analysis on the CGD platform with the whole human SR protein kinase 1 protein (SRPK1) (NCBI Reference: NP_003128.3) as query. This verified the annotation in the CGD and revealed no further homologs. Interestingly, except for C. glabrata, the most common pathogenic Candida spp. possess two SRPK homologs ( Supplementary Figure 1 ). The CAGL0F03905g-derived protein in C. glabrata CgSky1 has sequence identity of 60.35% to ScSky1 (NCBI BLASTp). This illustrates the phylogenetic relationships between the examined species: C. glabrata is more closely related to S. cerevisiae compared to the other Candida species that belong to the CTG clade (Muñoz et al., 2018). Thus, we decided to include CgSky1 in our further analyses to better understand the degree of evolutionary conservation of SR-kinases in Candida spp. Multiple sequence alignment showed that the domains that contain the ATP binding sites, the activation loop and the kinase-docking sites are particularly highly conserved between Homo sapiens, S. cerevisiae, C. glabrata, and C. albicans ( Figure 1 ). Interestingly, the CaSky2 protein has three insertions of amino acid strings of considerable length which are scattered throughout the kinase domain (alignment position 29–57, 94–108 and 153–320) that are not present in the other proteins ( Figure 1 ).

S. cerevisiae Sky1 has been characterized as a regulator of polyamine transport and ion homeostasis, as mutants lacking SKY1 are more resistant to toxic cations and polyamine concentrations (Erez and Kahana, 2001). Ion homeostasis is important for C. albicans and C. glabrata growth and proliferation, survival in the host, virulence mechanisms and resistance to antifungal drugs (Llopis-Torregrosa et al., 2016; Li et al., 2018). To test whether CgSky1, CaSky1, and CaSky2 are involved in similar cellular functions we generated C. glabrata mutant strains lacking SKY1 (Cgsky1Δ) and C. albicans strains lacking both SKY1 and SKY2 (Casky1Δ Casky2Δ). In addition, we used available C. albicans Casky1Δ and Casky2Δ (orf19.35) mutants (Ramírez-Zavala et al., 2022). As a control we used the S. cerevisiae BY4741 wild-type strain and a sky1Δ mutant (Scsky1Δ). In accordance with previous observations, the Scsky1Δ mutant was more resistant to osmotic stress caused by NaCl or LiCl and to high concentrations of the polyamine spermine (Erez and Kahana, 2001; Forment et al., 2002). Similar phenotypes were observed for C. glabrata mutants lacking SKY1 ( Figure 2 ). Deletion of SKY1 in C. albicans resulted in high resistance to LiCl and spermine, whereas lack of SKY2 rendered the cells partially resistant to these stressors. The Casky1Δ Casky2Δ double mutant had the same phenotype as the Casky1Δ deletion mutant. None of the tested C. albicans SKY deletion mutant strains exhibited altered resistance to NaCl, in contrast to the Scsky1Δ and Cgsky1Δ mutants ( Figure 2 ). Altogether, these results demonstrate a functional conservation of Sky1 among the examined species.

Since we found divergences in the protein sequence and phenotypic differences between the two C. albicans SRPKs CaSky1 and CaSky2, we decided to apply a high-throughput phenotypic screen to further characterize their cellular roles. Given that the two independently generated Casky1Δ and Casky2Δ mutant strains showed the same phenotypes, we utilized only strain A of the respective mutants for this analysis. The Casky1Δ and Casky2Δ mutant strains were screened for metabolic activity on 904 different nutrients and supplements and compared to the metabolic activity of the wild-type strain SC5314. Overall, there were fewer and less pronounced phenotypic differences between the Casky1Δ mutant and the wild type compared to the Casky2Δ mutant vs. wild-type strain SC5314 (data not shown). Notably, the Casky2Δ mutant had impaired growth on multiple dipeptide combinations as the sole nitrogen source ( Figure 3 ; Supplementary Figure 2 ). The Casky2Δ mutant was able to utilize only 104 out of the 268 tested dipeptides (38.8%), whereas the wild-type strain and the Casky1Δ mutant were able to utilize 146 (54.5%) and 125 (46.6%) dipeptides, respectively. Interestingly, the Casky2Δ mutant was as capable as the wild type in utilization of tripeptides, showing that the dipeptide growth defect is likely to be potentially specific ( Figure 3 ; Supplementary Figure 2 ).

To validate the observations from the phenotypic screen we performed growth curve analyses using defined medium and added the B strains of the Casky1Δ and Casky2Δ mutants and the corresponding complemented strains. Based on the results of the phenotypic screening we selected as the sole nitrogen source several dipeptides on which the Casky2Δ mutant showed a growth defect ( Figure 3 ): alanine-phenylalanine (Ala-Phe), alanine-tyrosine (Ala-Tyr) and valine-alanine (Val-Ala). Growth curve analyses confirmed that the Casky2Δ mutants exhibit significantly reduced growth compared to the wild-type strain and the Casky1Δ mutants on these particular dipeptides ( Figure 4 ). Interestingly, Casky1Δ grew better on Ala-Tyr as the sole nitrogen source compared to the wild-type strain ( Figure 4 ). We also tested growth on phenylalanine-serine (Phe-Ser), a dipeptide where both Casky1Δ and Casky2Δ mutants had impaired growth in the phenotypic screen. A similar defect was observed in the growth curve assay ( Figure 4 ). Further, both Casky1Δ and Casky2Δ mutants showed reduced growth on Phe-Ser compared to the wild type, with Casky2Δ mutant strains growing significantly worse than the Casky1Δ mutants. The SKY2 complemented strains either partially rescued the Casky2Δ mutant phenotype (Ala-Phe, Val-Ala) or exhibited growth comparable to the Casky2Δ mutant (Ala-Tyr, Phe-Ser). Both complemented strains were generated by re-introduction of allele A of SKY2. Comparison of the two SKY2 alleles revealed some differences in their sequences, which prompted us to test the growth of the heterozygous SKY2/sky2 mutant strains in which either allele A or allele B was still present on the selected dipeptides. Both heterozygous mutants showed comparable growth to the homozygous Casky2Δ mutant strains, indicating a gene dosage effect in which one SKY2 allele is not sufficient to complement the phenotype of the wild-type strain ( Supplementary Figure 3 ). Taken together, these results confirm the findings from the phenotypic screen that C. albicans Sky2 is required for assimilation of dipeptides as a nitrogen source.

We next performed a phosphoproteome analysis to identify the potential substrates responsible for the functional differences between CaSky1 and CaSky2. In brief, we grew the strains in YPD medium, extracted the proteins, and performed an LC-MS/MS analysis of tryptic peptides. TiO₂/ZrO₂-mediated phosphopeptide enrichment facilitated the identification and quantification of phosphopeptides with serine, threonine, and/or tyrosine phosphorylation. Phosphopeptide abundances were corrected against the corresponding master protein abundances (obtained from the non-enriched fraction) to enable a site-specific quantification. With this approach, we were able to identify 3946 proteins, 1663 phosphoproteins, 7243 phosphopeptides and 7727 phosphosites (82.05% serine, 16.57% threonine, 1.38% tyrosine) in total. Comparison of the fold change between the Casky1Δ mutant versus wild type and the Casky2Δ mutant versus wild type of all 3946 identified proteins revealed large differences in their quantile distributions (data not shown). Further, we identified 28 proteins significantly more abundant (log₂ ratios > 2) and 31 proteins less abundant (log₂ ratios < -2) in the Casky1Δ mutant versus wild type. On the other hand, we identified 31 proteins significantly more abundant (log₂ ratios > 2) and 142 proteins less abundant (log₂ ratios < -2) in the Casky2Δ mutant versus wild type, revealing differences between CaSky1 and CaSky2. The differences become especially clear on the phosphopeptide level when comparing the quantile distribution of all identified phosphopeptides (n = 7243) ( Figure 5A ) and the phosphopeptide abundance log₂ ratios of the Casky1Δ mutant vs. wild type and the Casky2Δ mutant vs. wild type ( Figures 5B, C ). For the Casky1Δ mutant we identified 268 phosphopeptides assigned to 224 phosphoproteins with significantly altered abundance (175 more abundant (log₂ ratios > 2), 49 less abundant (log₂ ratios < -2) compared to the wild-type strain SC5314 ( Figure 5B ; Supplementary Table 2 ). In contrast, we identified 237 phosphopeptides assigned to 210 phosphoproteins significantly altered in abundance (23 more abundant (log₂ ratios > 2), 187 less abundant (log₂ ratios < -2)) in the Casky2Δ mutant compared to the wild-type strain ( Figure 5C ; Supplementary Table 3 ).

To find out in which biological processes the identified proteins are involved, we applied an in-depth GO term analysis. All identified phosphoproteins that were significantly more abundant in the Casky1Δ mutant compared to the wild-type strain SC5314 belong to several different GO terms like cellular component organization, cell cycle processes, cytoskeleton organization, cellular response to stimulus and regulation of transcription by RNA polymerase II ( Figure 6A ). For the identified 49 significantly less abundant phosphoproteins in the Casky1Δ mutant compared to the wild type, positive regulation of protein kinase activity was the only significant (-log₁₀ p-values > 1.3) GO term that appeared in our analysis ( Figure 6A ). Although we could not identify any significant GO term for the 23 more abundant phosphoproteins in the Casky2Δ mutant compared to the wild type, 22 GO terms were identified for the 187 significantly less abundant phosphoproteins in the Casky2Δ mutant compared to the wild type, e.g. regulation of biological and cellular processes, regulation of metabolic processes, growth and response to starvation ( Figure 6B ).

Of the 49 less abundant phosphoproteins identified in the Casky1Δ mutant compared to the wild type, Hrk1 was the only protein kinase, with > 63 times (log₄ fold change) lower phosphopeptide abundance and no detected phosphorylation at S81 and S95 ( Supplementary Table 2 ). Since HRK1 encodes an uncharacterized protein kinase to which we had available homozygous mutants (independently derived mutants A and B), we tested their resistance to osmotic stress and toxic polyamine concentrations. Interestingly, hrk1Δ mutants exhibited increased resistance to high LiCl and spermine concentrations, similar to Casky1Δ mutants, suggesting that Hrk1 is either a direct target of CaSky1 or an indirect target acting in the same pathway ( Figure 7 ).

Among the 187 phosphoproteins that were significantly less abundant in the Casky2Δ mutant compared to the wild type, the transcription factors Fcr1 and Sfl1 as well as several protein kinases (Cst20, Gin4, Hsl1, Kic1, Ptk2, Sak1, Sat4, Swe1 and orf19.846) were identified as potential Sky2 targets. This suggests that CaSky2 is either part of a regulatory protein kinase network or that deletion of SKY2 affects several pathways in the cell. Another potential target was the di- and tripeptide transporter Ptr22. The phosphopeptides were 70–85 times (log₄ fold change) less abundant and no phosphorylation at S2, T3, and S39 was detected in the Casky2Δ mutant compared to the wild-type strain ( Supplementary Table 3 ). We found this particularly interesting since Ptr22 and Ptr2 are the only known dipeptide transporters in C. albicans, with Ptr22 having a broader substrate spectrum than Ptr2 (Dunkel et al., 2013). Thus, in addition to altered phosphorylation, the observed strongly decreased abundance of Ptr22 may also explain the growth defect of the Casky2Δ mutants on dipeptides as the sole nitrogen source ( Figures 3 , 4 ).

To test whether Sky2 regulates dipeptide utilization via the di- and tripeptide transporter Ptr22, we generated strains overexpressing PTR22 in the wild-type strain SC5314 and in the Casky2Δ mutant background and monitored their growth on the dipeptides Ala-Phe, Ala-Tyr, Phe-Ser, and Val-Ala as the sole nitrogen source. We also included two independently generated ptr22Δ mutants (strain A and B) as control, shown to have a growth defect on dipeptides (Dunkel et al., 2013). As expected, the two strains lacking PTR22 were unable to grow on the tested dipeptides ( Figure 8 ). Overexpression of PTR22 under control of the constitutive ADH1 promoter rescued the growth defect of the Casky2Δ mutants and significantly improved the growth of the wild-type strain on all dipeptides tested ( Figure 8 ).

It has been shown that C. albicans strains with defective uptake of di- and tripeptides are resistant to nikkomycins, polyoxins, and bacilysin, nucleoside antifungal agents that act as competitive inhibitors of chitin biosynthesis (Mehta et al., 1984; Payne and Shallow, 1985). Nikkomycin Z is in phase 2 clinical trial for treatment of coccidioidomycosis in humans (Nix et al., 2009) and we tested the susceptibility of the wild type and the mutant strains to this antifungal as well as to Polyoxin D. While these antifungals significantly impaired the growth of the wild-type strain, the ptr22Δ mutants were only slightly affected ( Figure 9 ). Strains lacking SKY2 exhibited resistance to both antifungals, but at a lower level than the ptr22Δ mutant strains. Growth of strains overexpressing PTR22 in the wild type or in the Casky2Δ mutant background was completely inhibited by both antifungals ( Figure 9 ). These results show that Ptr22 mediates the uptake of Nikkomycin Z or Polyoxin D, a function that is regulated in part by CaSky2.