Animal care and research were approved after deliberation by the Institutional Animal Care and Use Committee of the Experimental Animal Center at Jeonbuk National University. (Approval number: JBNU 2022-092). All experiments followed the experimental ethics guidelines. Animal experiments were conducted at the Core Facility Center for Zoonosis Research (Jeonbuk National University, South Korea).

Candida auris strains used in this study are listed in Table S1 in the Supplemental Material . The parental wild-type strain, B8441 (AR0387), was obtained from the Centers for Disease Control and Prevention (CDC). These isolates and the constructed mutant strains were stored as frozen stocks in 20% glycerol at -80°C until further use. Yeast strains were routinely cultured on YPD agar plates (2% agar in YPD broth: 1% yeast extract, 2% peptone, and 2% D-glucose) at 30°C for 24-48 hours. For liquid cultures, cells were grown in YPD broth at 30°C with shaking at 200 rpm. For experimental assays, cells were inoculated into fresh YPD broth and grown to mid-log phase (an optical density at 600 nm (OD₆₀₀) of 0.8) before being subjected to various treatments.

To generate gene deletion mutants, we used the nourseothricin resistance marker (CaNAT) flanked by 0.3- to 0.7-kb 5′ and 3′ regions of each target gene, including SAPA1, SAPA2, SAPA3, SAPA4, SAPA5, SAPA6, and SAPA7. Each gene disruption cassette containing a selection marker was constructed using double-joint PCR. To amplify the flanking regions of a target gene, we used L1-L2, and R1-R2 primer pairs in the first round of PCR. The CaNAT selection marker was amplified by PCR using the plasmid pV1025 containing the CaNAT gene as a template and the primer pairs listed in Table S1 in the Supplemental Material . The first round of PCR products of the flanking regions and CaNAT marker were purified together and used as templates for the second round of double-joint PCR. In the second round of PCR, 5′- and 3′- gene disruption cassettes containing split CaNAT selection markers were amplified by L1-split primer 2 and R2-split primer 1, respectively ( Table S1 in the Supplemental Material ).

For the transformation of C. auris with gene disruption cassettes, we used a lithium acetate/heat-shock protocol with modifications. Cells were cultured overnight at 30°C in 50 mL YPD broth with shaking. We centrifuged 1.2 mL of cultured cells, washed them with dH₂O and lithium acetate buffer (100 mM lithium acetate, 10 mM Tris, 1 mM EDTA, pH 7.5), and resuspended them in 300 μL of lithium acetate buffer. The transformation was set up with 10 μL of denatured salmon sperm DNA (Sigma, cat no. D9156), 100 μL of the competent cells, 500 μL of 50% PEG4000 (Sigma, cat no. P4338), and 50 μL of the amplified gene deletion cassette. The transformation mixture was incubated at 30°C for 6 hours with intermittent vortexing. Subsequently, the cells were subjected to a 20-minute heat shock at 42°C followed by 1 minute of cooling on ice. The cells were then pelleted, resuspended in 1 mL of YPD medium, and incubated at 30°C for 1 hour with shaking. After the incubation, the cells were washed twice with fresh liquid YPD medium and then spread onto selective YPD agar plates supplemented with 600 µg/mL nourseothricin. The plates were then incubated at 37°C for 2 days. We confirmed the desired genotype of each positive nourseothricin-resistant transformant by diagnostic PCR and Southern blot ( Figure S1 ).

To confirm the phenotypes of the sapa3Δ mutant, we constructed corresponding complemented strains, in which each wild-type allele was re-integrated into its native locus. To generate each full-length gene fragment, Phusion-PCR was performed using genomic DNA from the wild-type B8441 strain as a template and each primer pair listed in Table S1 in the Supplemental Material . The resulting fragments were directly cloned into the TOPO vector (Invitrogen) to generate the plasmids pTOP-SAPA3. After confirming the target sequence, the CaHYG inserts were sub-cloned into each pTOP vector to produce the pTOP-SAPA3-HYG. For the targeted re-integration into its native locus, pTOP-SAPA3-HYG was linearized by StuI, and introduced into each mutant by the lithium acetate heat-shock method. The correct genotype of the complemented strain was confirmed by diagnostic PCR ( Figure S1 ).

SAP activity was tested using the YCB-BSA method with slight modifications. C. auris strains were cultured overnight in 2 mL YPD broth at 30°C, washed, and resuspended in 1 mL of dH₂O. Next, 3 μL of the suspended cells were then spotted on YCB-BSA plates (containing 23.4g yeast carbon base per liter and 0.2% BSA) and incubated at different temperatures for 3 days. The thickness of the halo was measured to determine the SAP activity, and the experiment was biologically repeated three times.

Cells of C. auris wild-type and mutant strains were cultured overnight at 30°C in 2 mL YPD liquid medium, washed twice with H₂O, and then resuspended in MOPS-buffered RPMI-1640 media (pH 7.4 with 0.165 M MOPS and 2% glucose). For the crystal violet assay, the cell suspension was prepared with a concentration equivalent to an OD₆₀₀ of 0.5. Subsequently, 200 μL of the cell suspension was loaded into each well of a 96-well plate, each inhibitor was treated at a concentration of 10 μM, and the cultures were incubated at 37°C and 220 rpm for 1 day. The next day, the cell suspension was completely removed, and the samples were completely dried in a dry oven at 65°C. To each well, 150 μL of a 0.3% crystal violet solution was added, and the samples were stained at room temperature for 10 minutes. Subsequently, the crystal violet solution was completely removed, and the wells were washed three times with ddH₂O. The samples were then dried in a dry oven at 65°C. Next, 200 μL of 33% acetic acid was added to each well, and the crystal violet was dissolved completely for 1 minute. The solution was transferred to a clean 96-well plate, and the absorbance was read at OD₅₉₅.

Cells were cultured overnight at 30°C in YPD medium and subcultured to OD₆₀₀ of 0.8 in fresh YPD medium. After sub-culturing the cells were suspended in YPD broth supplemented with 100 mM HU and incubated at 30°C for 24 hours. After hyphal growth, each sample was fixed using a 10% formalin solution and stained with a calcofluor white solution. The samples that had been fixed and stained were then incubated in a dark environment for 30 minutes before being photographed.

To analyze the growth and sensitivity to various stresses of WT and mutant strains. To do this, C. auris cells grown overnight at 30°C were serially diluted tenfold, four times (final dilution 1:104) and spotted onto YPD plates. For growth measurements, plates were incubated at 30°C, 37°C, and 42°C. Growth was assessed qualitatively by photographing the plates 1 day later.

Various stresses were imposed by adding to the media chemical agents that would impose stress on the cells. Osmotic stress was supplied as sorbitol. Cation and salt stress were imposed with NaCl or KCl. Oxidative stress was supplied with hydrogen peroxide, tert-butyl hydroperoxide (an organic peroxide), menadione (a superoxide anion generator), or diamide (a thiol-specific oxidant). Membrane destabilizing stress was imposed with SDS and cell-wall destabilizing stress was imposed with CR, fluconazole, amphotericin B, or caspofungin for antifungal drug susceptibility. Cells were incubated at 30°C and photographed 1 to 4 days after treatment.

To evaluate the virulence of C. auris wild-type and Ras/cAMP/PKA mutant strains in vivo, we used a murine systemic infection model with reference to previous studies (Gao et al., 2021; Kim et al., 2021). Fungal cells were incubated overnight at 30°C in YPD broth and washed three times with PBS. Cell concentrations were measured and adjusted to 10⁸ cells/mL in PBS. To confirm colony-forming units (CFUs) and viability of the inoculum, the diluted cells were plated onto YPD agar plates and incubated at 37°C for 24 h. SPF/VAF-confirmed inbred 6-week-old female mice of the BALB/c (AnNCrlOri) strain were used for this study (ORIENT BIO INC., South Korea), and they were habituated for one week before the experiment. For infection, the mice were restrained, and their tails were placed in warm (40°C) water to expand the lateral veins. The 100 µL of cell suspension was injected intravenously. Daily monitoring of survival was performed and abnormal behavior (head tilt or body spinning) was judged as a symptom of infection (Gao et al., 2021), and the mice were sacrificed as a humane endpoint for the experiment. The survival curves were analyzed using the Log-rank (Mantel-Cox).

Total RNA was extracted from C. auris wild-type and Ras/cAMP/PKA mutant strains cultured overnight at 30°C in YPD broth. Briefly, cells were collected by centrifugation after reaching an OD₆₀₀ of 0.8, frozen in liquid nitrogen, and lyophilized. For stress conditions, 10 mL of the culture was sampled for the basal state, and the remaining 30 mL was further incubated with stress agents. Total RNA was isolated by the Trizol extraction method with Easy-blue (Intron). Complementary DNA (cDNA) was synthesized from purified total RNA using reverse transcriptase (Thermo Scientific). Quantitative PCR was performed using specific primer pairs for each gene and the CFX96TM Real-Time system (Bio-Rad). ACT1 expression was used for normalization. Statistical analysis was performed using one-way ANOVA, followed by Bonferroni’s multiple-comparison test. All experiments were conducted in triplicate and repeated thrice biologically.

To elucidate the role of the Ras/cAMP/PKA pathway in modulating SAP activity in C. auris, we measured SAP activity in mutant strains where the Ras/cAMP/PKA pathway was inactivated. We assessed SAP activity in wild-type and mutant strains at 30°C, 37°C, and 42°C ( Figures 1A–C ). Compared to the wild-type strain, tpk1Δ tpk2Δ, cdc25Δ, and cyr1Δ strains exhibited a noticeable reduction in SAP activity at 30°C ( Figure 1A ). This reduction was evident in the measurements of average halo thickness, with tpk1Δ tpk2Δ, cdc25Δ, and cyr1Δ showing thicknesses of 0.118 cm, 0.405 cm, 0.401 cm respectively, compared to the wild-type’s 0.538 cm ( Figure 1A ). The tpk1Δ tpk2Δ strain, in particular, displayed a severe decrease in SAP activity across all temperature conditions. As the temperature increased, SAP activity in the cdc25Δ and cyr1Δ strains reached levels closer to the wild-type ( Figures 1B, C ). However, the tpk1Δ tpk2Δ strain consistently showed significantly lower SAP activity than the wild-type strain, even at higher temperatures. At 37°C, the average halo thickness of the wild-type strain was 0.639 cm, while tpk1Δ tpk2Δ exhibited a thickness of 0.313 cm ( Figure 1B ). At 42°C, the wild-type’s average halo thickness was 0.721 cm, whereas tpk1Δ tpk2Δ showed a thickness of 0.356 cm ( Figure 1C ).

Our findings indicate that like in C. albicans, the SAP activity in C. auris is temperature-dependent, providing additional evidence that the Ras/cAMP/PKA pathway regulates this activity. Notably, our data support the pivotal role of PKA catalytic subunits in modulating SAP activity in C. auris. However, the decreased dependence of the Ras/cAMP/PKA signaling pathway in regulating SAP activity with increasing temperature suggests the involvement of additional signaling pathways in SAP regulation. In C. albicans, the MAPK pathway, in tandem with the Ras/cAMP/PKA signaling pathway, modulates SAP gene expression (Naglik et al., 2004).

To identify SAP-encoding genes in C. auris, we leveraged the Candida Genome Database, specifically looking for proteins containing the aspartic peptidase domain (IPR033876). Our search revealed that while C. albicans has 10 proteins with this domain, C. auris features 14 proteins. Phylogenetic analysis showed a notable divergence between the SAPs of C. auris and C. albicans ( Figure 2A ). A BLAST analysis performed on C. albicans SAPs against the C. auris genome revealed that Sap1, Sap2, Sap3, Sap4, Sap5, and Sap6 primarily aligned with B9J08_001518. Similarly, Sap7, Sap9, and Sap10 were most closely associated with B9J08_001958, and Sap8 first aligned with B9J08_001534 ( Table S2 ). When we performed the reverse BLAST analysis using C. auris SAPs against the C. albicans genome, we found that B9J08_000398, B9J08_001958, B9J08_003911, B9J08_003912, and B9J08_005335 were best aligned with Sap9. Similarly, B9J08_001546, B9J08_004629, and B9J08_004888 primarily matched with Sap8. Furthermore, each of B9J08_005287, B9J08_001534, B9J08_005421, B9J08_001518, B9J08_002149, and B9J08_005371 demonstrated a first match with Sap5, Sap4, Sap3, Sap2, Yps7, and Arp1, respectively ( Table S2 ).

Due to the overall low similarity between the SAPs of C. auris and C. albicans and the lack of one-to-one protein matches in the BLAST analysis, we decided to focus our investigations on the seven proteins with high scores and low E-values in the BLAST analysis of C. albicans SAPs against the C. auris genome. These proteins, labeled as Sapa1 to Sapa7 in C. auris (B9J08_001518: Sapa1, B9J08_001534: Sapa2, B9J08_001546: Sapa3, B9J08_004629: Sapa4, B9J08_005421: Sapa5, B9J08_001958: Sapa6, B9J08_000398: Sapa7), were selected for further analysis ( Figures 2A, B ). This outcome underscores the significant inter-species divergence within the aspartic peptidase domain.

To elucidate the role of SAPs in C. auris, we generated knockout mutants for each of the seven SAP genes ( Figure S1 ). Subsequently, we measured and compared the SAP activity of these mutants (sapa1Δ, sapa2Δ, sapa3Δ, sapa4Δ, sapa5Δ, sapa6Δ, and sapa7Δ) with the wild-type strain at 30°C, 37°C, and 42°C ( Figures 3A–C ). At 30°C, the wild-type strain demonstrated an average halo thickness of 0.538 cm. The sapa1Δ, sapa2Δ, sapa4Δ, sapa5Δ, sapa6Δ, and sapa7Δ mutants, with respective average halo thicknesses of 0.493 cm, 0.569 cm, 0.539 cm, 0.567 cm, 0.514 cm, and 0.558 cm, showed no significant deviation from the wild-type strain ( Figure 3A ). However, the sapa3Δ mutant manifested a pronounced decrease in SAP activity, with an average halo thickness of 0.206 cm, signifying a substantial reduction of about 61% ( Figure 3A ).

This pattern persisted across all temperature conditions. As the temperature rose, the SAP activity of the wild-type strain and all mutant strains, except sapa3Δ, escalated by roughly 30-40% compared to measurements at 30°C ( Figures 3B, C ). Yet, the sapa3Δ exhibited only a minimal increase in SAP activity. At 37°C, the wild-type strain displayed an average halo thickness of 0.672 cm, while sapa3Δ showed an average halo thickness of 0.269 cm − about 59% less than the wild-type ( Figure 3B ). Similarly, at 42°C, the wild-type registered an average halo thickness of 0.725 cm, while sapa3Δ showed a notable drop in SAP activity, with an average halo thickness of 0.245 cm, signifying a 66% decrease compared to the wild-type strain ( Figure 3C ).

To confirm the role of Sapa3, we generated the sapa3Δ::SAPA3 complemented strain ( Figure S1 ), and measured its SAP activity at 30°C, 37°C, and 42°C compared to the wild-type and sapa3Δ strains ( Figure 3D ). The complemented strain exhibited SAP activity levels equivalent to those of the wild-type strain ( Figure 3D ), affirming that SAP3 encodes a major SAP in C. auris and that disruption of Sapa3 results in a loss of SAP activity irrespective of temperature. All these data emphasize the critical role of Sapa3 as the primary SAP in C. auris.

Our findings highlight the regulatory role of the Ras/cAMP/PKA pathway on SAP activity and identify Sapa3 as the major SAP in C. auris. Subsequently, we investigated whether the Ras/cAMP/PKA pathway directly modulates Sapa3. At 30°C, inactivation of the Ras/cAMP/PKA pathway resulted in a significant decline in SAPA3 expression ( Figure 4A ). Specifically, the cyr1Δ, cdc25Δ, ras1Δ, and tpk1Δ tpk2Δ strains displayed around 60-80% decrease in SAPA3 expression compared to the wild-type strain ( Figure 4A ). As the temperature increased, however, the impact of the Ras/cAMP/PKA inactivation on SAPA3 expression became less apparent. At 37°C, we observed a significant reduction in SAPA3 expression in the cdc25Δ and tpk1Δ tpk2Δ strains, while the cyr1Δ and ras1Δ strains exhibited similar SAPA3 expression levels to the wild-type strain ( Figure 4B ). At 42°C, the inactivation of the Ras/cAMP/PKA pathway did not lead to a decrease in SAPA3 expression ( Figure 4C ). These results suggest that although SAPA3 is a key SAP whose expression is modulated by the Ras/cAMP/PKA pathway, this regulation appears to diminish at higher temperatures. Notably, even at 42°C, the tpk1Δ tpk2Δ strain displayed decreased SAP activity. However, this was not coupled with a significant reduction in SAPA3 expression compared to the wild-type strain, indicating the potential influence of other SAP genes on SAP activity in C. auris.

We next conducted phenotypic analyses to evaluate the role of SAP genes in biofilm formation, filamentous growth, and stress resistance in C. auris. Given that in C. albicans, SAP genes are implicated in biofilm development and pseudohyphae formation, we sought to ascertain if similar functionalities exist in C. auris.

Biofilm formation was gauged using the crystal violet assay. The biofilm-forming potential of the sapa2Δ mutant was marginally diminished by about 14% relative to the wild-type, though the difference did not reach statistical significance ( Figure 5 ). On the other hand, the sapa3Δ mutant exhibited a 28% decrease in biofilm formation ( Figure 5 ). Notably, the sapa6Δ mutant showed an increase in biofilm formation by around 14% compared to the wild-type ( Figure 5 ). These observations underscore the significance of SAP genes in C. auris biofilm formation, especially spotlighting the integral function of Sapa3.

To probe into the pseudohyphae formation capability, the wild-type and mutant strains were incubated in 100 mM hydroxyurea over a 24-hour period. The outcome highlighted that all strains, including the wild-type, manifested standard pseudohyphae formation ( Figure S2 ), suggesting that SAP genes are not critical for filamentous growth in C. auris. We also assessed the resistance of SAP gene mutants to a range of stress conditions. The wild-type and mutant strains exhibit similar growth patterns at 30°C, 37°C, and 42°C ( Figure S3 ). Additionally, with the exception of a slightly diminished resistance of sapa5Δ to diamide and sapa6Δ to DTT, we observed no significant differences in resistance between wild-type and SAP gene mutants when exposed to various antifungal drugs, oxidative, osmotic, genotoxic, cell wall and membrane stresses, heavy metal challenges, ER stress, and acidic pH conditions ( Figure S3 ). These results collectively indicate that SAP genes have a limited influence on the stress defense mechanisms in C. auris.

Next, we explored the contribution of Sapa3 to C. auris pathogenicity using and BALB/c mouse model. The mice were intravenously infected with the wild-type, sapa3Δ, and sapa3Δ::SAPA3 strains, and their survival rate was monitored over 36 days. Compared to the wild-type and complemented strains, the sapa3Δ mutant manifested a significant decrease in pathogenicity ( Figure 6 ). While all mice infected with the wild-type and complemented strains succumbed within 15 and 22 days, respectively ( Figure 6 ), a portion of mice infected with the sapa3Δ mutant survived until day 36 ( Figure 6 ). These results underscore the pivotal role of SAP as a key virulence factor in C. auris.