2.2. The Effect of SFP1 Deletion on Cell Susceptibility to LL-37 Is Cell Wall-Related

The cell wall is a dynamic structure in response to environmental changes and various stress conditions, such as the presence of human AMPs [14,15,22]. Since deletion of SFP1 leads to reduced LL-37 susceptibility (Figure 1A,B), cell susceptibility to tunicamycin was also examined to correlate Sfp1 and the cell wall. Tunicamycin is a nucleoside antibiotic that affects the cell wall by inhibiting N-linked glycosylation and significantly inhibits C. albicans biofilm formation and maintenance [44]. As shown in Figure 2A, the sfp1Δ/sfp1Δ mutant was much more resistant to tunicamycin than the wild-type and SFP1-reintegrated strains.

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Figure 2

Assessment of the connection of Sfp1 with cell wall integrity and cellular response to LL-37. (A) Cells were serially diluted tenfold, spotted onto SC agar plates with or without tunicamycin (3 and 5 μg/mL), and incubated at 30 °C for 7 days. Representative data from at least three experiments with identified results are shown. WT: wild type; sfp1Δ/sfp1Δ: the SFP1 deletion mutant; sfp1Δ::SFP1/sfp1Δ::SFP1: the SFP1-reintegrated strain. CFU: colony forming unit. (B) Cells were pretreated with different concentrations of LL-37 for 30 min, serially diluted tenfold and spotted onto SC agar plates with or without 30 μg/mL congo red and 180 μg/mL calcofluor white. The agar plates were incubated at 30 °C for 5 days. Representative data from at least three experiments with identified results are shown. (C) Mkc1 activation induced by LL-37 in different C. albicans strains. Cells were treated with or without LL-37 (8 μg/mL) or tunicamycin (5 μg/mL) for 30 min. Equal amounts of protein extracts from each sample were loaded. p-Mkc1 was monitored by Western blotting and analyzed with ImageJ software. The Act1 band of each sample served as the loading control and was used to normalize the phosphor-Mkc1 levels indicated by the fold change values. The data are representative of three independent experiments with identical results.

To further determine the relationship among Sfp1, the cell wall and the cell response to LL-37, the susceptibility of cells pretreated with or without LL-37 to the cell wall-disrupting agents Calcofluor white (CFW) and Congo red (CR) and activation of the Mkc1 MAP kinase were also assessed. CFW and CR interact with chitin and β-glucan and disrupt cell wall biosynthesis [45]. In the spot assays, with treatment at different concentrations of LL-37, the sfp1Δ/sfp1Δ mutant was overall more resistant to CFW and CR than the wild-type and SFP1-reintegrated strains (Figure 2B). Moreover, the C. albicans Mkc1 signaling pathway plays an important role in cell wall construction and is activated upon cell wall stress, such as that caused by tunicamycin [46]. Tunicamycin was thus used as a positive control to detect Mkc1 activation by phosphorylation in the Western blot analysis. As expected, treatment with tunicamycin activated Mkc1 in all of the tested strains (Figure 2C). In addition, increased levels of phosphorylated Mkc1 (p-Mkc1) were detected in the wild-type and SFP1-reintegrated strains with LL-37 treatment compared to those without treatment. However, the sfp1Δ/sfp1Δ mutant displayed elevated levels of p-Mkc1, not only in cells treated with LL-37 but also in cells not treated (Figure 2C). A graph integrating the quantification of three independent Western blot analysis was also provided in Figure S1 (in the Supplementary Materials). Together, the results suggest that LL-37 is associated with activation of the Mkc1 signaling pathway, and the cell response to LL-37 is related to the impact of SFP1 deletion on cell wall integrity.

2.3. Deletion of SFP1 Attenuates HAC1 mRNA Splicing and UPR-Responsive Gene Activation upon LL-37 Treatment

In C. albicans, the ER stress sensor Ire1 affects cell wall integrity, and its downstream effector Hac1 regulates the expression of genes related to cell wall biosynthesis [38,39]. Interestingly, as demonstrated in Figure 1 and Figure 2, LL-37 induces the Mkc1-mediated cell wall integrity pathway, and the impacts of Sfp1 on the cell response to LL-37 are cell wall-associated. These results thus allowed us to consider a possible correlation between the ER stress response and Sfp1 when cells were treated with LL-37.

The UPR is a cellular response related to ER stress and is characterized by the splicing of HAC1 mRNA [25]. To link the UPR and Sfp1, reverse transcription PCR was performed to detect spliced HAC1 mRNA. In this study, cells treated with tunicamycin, which is also known to induce ER stress and activate the UPR, were included as positive controls. In Figure 3, similar to tunicamycin, LL-37 activated the UPR, which is indicated by the elevated levels of spliced HAC1 mRNA in the wild-type and SFP1-reintegrated strains compared to those without LL-37 treatment. However, HAC1 mRNA was poorly spliced not only in the sfp1Δ/sfp1Δ mutant without LL-37 treatment but also in that with treatment (Figure 3). Moreover, the levels of spliced HAC1 mRNA in the sfp1Δ/sfp1Δ mutant appeared to be lower than those in the wild-type and SFP1-reintegrated strains (Figure 3). This result is somehow consistent with the finding that the sfp1Δ/sfp1Δ mutant is more resistant to tunicamycin than the control cells (Figure 2A).

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Figure 3

Detection of HAC1 mRNA splicing. Cells were treated with or without LL-37 (8 μg/mL) or tunicamycin (5 μg/mL) for 30 min. Total RNA was extracted, and RT–PCR was performed. U: unspliced HAC1; S: spliced HAC1. The size difference between the unspliced and spliced cDNA is 19 bp. The data are representative of three independent experiments with identical results. M: DNA marker.

In response to ER stress, C. albicans HAC1 mRNA is spliced to produce the Hac1 protein, which then activates the transcription of target genes [39]. To further connect UPR signaling and Sfp1, the expression of several UPR-responsive genes, PMT4, YSY6, GAA1 and ERD2 [47], under stress conditions was determined by real-time quantitative PCR (qPCR). Pmt4 is a protein mannosyltransferase, while Ysy6 is involved in the regulation of the UPR [48,49]. Moreover, Gaa1 is a putative subunit of the glycosylphosphatidylinositol (GPI)-anchor transamidase [50], and Erd2 is a protein possibly involved in secretion [51]. Transcription of these four genes was increased in the wild-type and SFP1-reintegrated strains with either LL-37 or tunicamycin treatment compared to that without treatment. However, the expression of these genes was not activated in the sfp1Δ/sfp1Δ mutant with LL-37 treatment compared to that without treatment (Figure 4). Together, these results indicate that LL-37 can induce ER stress and activate the UPR. In addition, deletion of SFP1 attenuates the UPR, suggesting that Sfp1 normally contributes to the stress response to maintain ER homeostasis.

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Figure 4

Effect of LL-37 and deletion of SFP1 on the expression of UPR genes. Cells were treated with or without LL-37 or tunicamycin for 30 min. Real-time quantitative PCR was used to detect UPR gene expression. The PMA1 transcripts were used as an endogenous control. The results are expressed as the mean ± SD of three independent assays. * p < 0.05; ** p < 0.01.

2.4. Deletion of SFP1 also Attenuates ROS-Induced Toxicity and Oxidation of Ero1 Promoted by LL-37

Reactive oxygen species (ROS) are natural byproducts of aerobic metabolism and critical in cell signaling and maintenance of cellular redox homeostasis [52]. However, excessive ROS accumulation can evoke cell toxicity and death. Recently, Sfp1 was found to regulate the oxidative stress response in C. albicans [43]. Moreover, the toxicity of cell wall stress toward C. albicans is mediated by ER-derived ROS [47]. Based on our findings described above, we proposed that LL-37 may alter the ER redox state and that Sfp1 is involved in this process.

The accumulation of intracellular ROS in the wild-type, sfp1Δ/sfp1Δ and SFP1-reintegrated strains with and without LL-37 treatment was compared (Figure 5A). Cells were stained with the nonpermanent ROS indicator 2′,7′-dichlorodihydrofluorescein diacetate (H₂DCFDA), which can be cleaved by intracellular esterases to convert into the highly fluorescent 2′,7’-dichlorofluorescein (DCF). The mean fluorescence intensity of DCF was proportional to ROS levels and determined by flow cytometry. As shown in Figure 5A, the fluorescence intensity was enhanced in the wild-type and SFP1-reintegrated strains treated with increasing concentrations of LL-37 compared to that without treatment. However, the levels of intracellular ROS in the sfp1Δ/sfp1Δ mutant showed no significant changes among cells with or without LL-37 treatment (Figure 5A). Moreover, ROS accumulation can cause oxidative stress and activate antioxidant gene expression. To further validate the involvement of Sfp1 in the regulation of oxidative stress induced by LL-37, the expression of the oxidative stress responsive (OSR) genes SOD1, CAT1 and GPX2 was also detected. The SOD1, CAT1 and GPX2 genes encode superoxide dismutase, catalase and glutathione peroxidase, respectively [43]. As shown in Figure 5B, the expression of these antioxidant genes was upregulated in all of the tested strains with LL-37 treatment compared to that without treatment. However, the sfp1Δ/sfp1Δ mutant exhibited much higher gene activation than the wild-type and SFP1-reintegrated strains (Figure 5B). Finally, to link ROS accumulation and cellular toxicity in all of the tested strains, lipid peroxidation products were measured using the thiobarbituric acid reactive substance (TBARS) assay. This assay detects malondialdehyde (MDA), which is a product generated from lipid oxidation of unsaturated fatty acids, and MDA is a common marker of oxidative stress [53]. In Figure 5C, the results showed that the levels of MDA were enhanced in the wild-type and SFP1-reintegrated strains treated with LL-37 compared to those without treatment. However, the MDA level of the sfp1Δ/sfp1Δ mutant did not change between cells treated with and without LL-37 and was overall lower than that in the wild-type and SFP1-reintegrated strains upon LL-37 treatment (Figure 5C).

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Figure 5

Oxidative stress in cells treated with or without LL-37. (A) To measure intracellular ROS accumulation, cells were treated with the indicated concentrations of LL-37 for 30 min, followed by incubation with 20 μg/mL H₂DCFDA. The mean fluorescence of 10,000 cells was determined by flow cytometry. The results are expressed as the mean ± SD of three independent assays. * p < 0.05; *** p < 0.001. (B) The expression of antioxidant genes was detected using real-time quantitative PCR. The PMA1 transcripts were used as an endogenous control. The results are expressed as the mean ± SD of three independent assays. * p < 0.05; ** p < 0.01. (C) Detection of lipid peroxidation as a marker of oxidative stress. Cells were treated with or without LL-37 (8 μg/mL) for 30 min, and the MDA level was measured. The results are expressed as the mean ± SD of three independent assays. * p < 0.05; ** p < 0.01.

Ero1 is an ER-resident oxidoreductase and is important for the formation of protein disulfide bonds in the ER [47,54,55]. Moreover, the reactivation of Ero1 by molecular oxygen is one of the causes of ROS generation [56,57]. In this process, the reduced (red) form Ero1 accepts the electrons from oxygen accompanying the generation of hydrogen peroxide. Then, the oxidized (ox) form of Ero1 catalyzes oxidative protein folding through disulfide exchanges [58,59]. In other words, ER redox homeostasis is established based on a balance between Ero1 activation and inactivation [60]. Since deletion of SFP1 diminished the effects of intracellular ROS accumulation and activation of the UPR pathway induced by LL-37 (Figure 3 and Figure 5A), these findings raise the possibility that Sfp1 affects ER redox homeostasis when cells cope with the stress induced by LL-37. To test this possibility, the redox state of Ero1 was determined using the thiol-reactive reagent 4-acetamido-4′-maleimidylstilbene-2,2′-disulfonic acid (AMS). AMS reacts with free cysteine thiols of Ero1 (red), but it cannot react with Ero1(ox) that contains no or low levels of free thiols. Therefore, after reacting with AMS, Ero1(red) should have a larger molecular weight than Ero1(ox) [47,58]. As shown in Figure 6, Western blotting was performed to detect Flag-tagged Ero1 using an anti-Flag antibody. Similar to tunicamycin treatment, LL-37 caused a 3- to 4-fold increase in the production of Ero1(ox) in both the wild-type and SFP1-reintegrated strains (Figure 6A,B). Conversely, there was no significant difference in the ratio of Ero1(ox)/Ero1(red) in the sfp1Δ/sfp1Δ mutant with LL-37 treatment compared to that without treatment (Figure 6).

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Figure 6

Effect of LL-37 and deletion of SFP1 on the redox state of Ero1. (A) After treatment with LL-37 or tunicamycin for 30 min, total proteins were harvested. The proteins were resolved by nonreducing 6% SDS–PAGE. Western blotting was performed to detect Ero1-Flag using an anti-Flag antibody. (B) The ratio of Ero1(ox)/Ero1(red) was obtained by analysis using ImageJ software. The results are expressed as the mean ± SD of three independent assays. * p < 0.05; ** p < 0.01.

In summary, LL-37 induces intracellular ROS accumulation, causes lipid peroxidation, an indicator of oxidative damage to cells, and enhances Ero1 oxidation. These results indicate that ER homeostasis is affected by LL-37. Moreover, these effects were observed in the wild-type and SFP1-reintegrated strains but not in the sfp1Δ/sfp1Δ mutant. Our findings further suggest that Sfp1 normally participates in the stress response to maintain ER homeostasis.

4.2. C. albicans Strains and Growth Conditions

The C. albicans strains used in this study are listed in Table S1 (in the Supplementary Materials). The cells were routinely grown in YPD medium (1% yeast extract, 2% peptone and 2% glucose) and synthetic complete (SC) medium (0.67% yeast nitrogen base [YNB] with ammonium sulfate, 2% glucose and 0.079% complete supplement mixture). Plates were prepared with 1.5% agar. One colony was inoculated into YPD broth and incubated at 30 °C overnight (~16 h) with shaking at 180 rpm. This culture was harvested by centrifugation, washed twice with sterile phosphate-buffered saline (PBS), subcultured in SC medium (with an initial concentration ~1.5 × 10⁷ cells/mL) and grown to the exponential phase for further experiments. For the induction of the MAL2 promoter and BSA degradation assay, YPM medium (1.0% yeast extract, 2.0% meat peptone and 2% maltose) and YCB-BSA-YE medium (1.17% Difco yeast carbon base, 0.01% BSA, 0.001% yeast extract, adjusted to pH 4.4 with HCl) were used separately as previously described [77].

4.3. Cell Susceptibility to LL-37 and Other Agents

Cell susceptibility to LL-37 was examined by CFU counting and PI staining. Briefly, C. albicans cells (~3.0 × 10⁷ cells/mL) were resuspended in LYM medium [78] containing different concentrations of LL-37 and incubated at 37 °C with slow shaking (100 rpm) for 30 min. Subsequently, cells were grown on YPD agar plates and incubated at 30 °C for 48 h, and the number of CFUs was counted. For PI staining, cells were treated with or without different concentrations of LL-37 for 30 min as described above and collected by centrifugation. After washing twice with PBS, cells were resuspended in PBS containing 5 μg/mL PI and incubated at room temperature for 5 min [43]. The fluorescence intensity of PI-positive cells was measured using an Accuri C6 flow cytometer (BD Biosciences, San Jose, CA, USA) with the FL2 filter. Cells without LL-37 treatment were used as negative controls. The average fluorescence intensity from at least three experiments was determined.

Cell susceptibility to cell wall-disrupting agents, CR and CFW, and the ER stress inducer tunicamycin was determined using a spot assay. Cells were treated with or without different concentrations of LL-37 and incubated at 37 °C for 30 min. Subsequently, cells were collected by centrifugation, washed twice with PBS and resuspended in sterile double-distilled water (ddH₂O). In this case, 10 microliters of each 10-fold serially diluted C. albicans cells were spotted onto SC agar plates containing CR (30 μg/mL), CFW (180 μg/mL) or different concentrations of tunicamycin. Cell viability was recorded after incubation at 30 °C for 1–7 days.

4.4. Assay for Mkc1 Phosphorylation

Protein extraction was performed as previously described [79,80] with some modifications. Briefly, cells were treated with or without LL-37 (8 μg/mL) and tunicamycin (5 μg/mL) for 30 min, harvested and washed twice with PBS. The cell pellets were suspended in lysis buffer (50 mM HEPES [pH 7.4], 5 mM ethylenediaminetetraacetic acid [EDTA], 1% Triton X-100, 1 mM NaF, 1 mM Na₃VO₄, 0.2 mM phenylmethylsulfonyl fluoride [PMSF], 4 μM leupeptin, 1 μM pepstatin A and 1 μg/mL aportinin) with acid-washed glass beads. The mixture was vortexed for 30 s and placed on ice for 30 s, and this process was repeated 10 times. Insoluble material was removed by centrifugation at 4 °C at 12,000× g for 10 min, and the supernatant of soluble proteins was collected. Protein concentration was measured using the Bradford Protein Assay (Bio–Rad, Hercules, CA, USA).

For Western blotting to detect phosphorylation of Mkc1, 30 μg of protein was mixed with 4x sample buffer (250 mM Tris-HCl [pH 6.8], 40% glycerol, 10% β-mercaptoethanol, 8% sodium dodecyl sulfate [SDS] and 0.4% bromophenol blue), boiled for 5 min and placed on ice for 5 min. Each protein sample was then separated by 10% SDS–PAGE and transferred onto a polyvinylidene difluoride (PVDF) membrane. The membrane was blocked overnight with 5% nonfat milk at room temperature. Anti-phospho-p44/p42 MAPK antibody (#4370; Cell Signaling Technology, Danvers, MA, USA) and anti-β-actin antibody (GTX109639; GeneTex, Hsinchu, Taiwan) were used to detect phosphor-Mkc1 and Act1, respectively. Horseradish peroxidase (HRP)-conjugated goat anti-rabbit IgG (GTX213110–01; GeneTex) was used as the secondary antibody. The blots were visualized using enhanced chemiluminescence reagents (PerkinElmer Inc., Waltham, MA, USA) and an ImageQuant LAS 4000 mini biomolecular imager (GE Health care, Chicago, IL, USA). Anti-β-actin was used as a loading control, and the bands were analyzed using ImageJ software [81].

4.5. Assay for HAC1 mRNA Splicing

To examine splicing of C. albicans HAC1 mRNA, cells were treated with or without LL-37 (8 μg/mL) and tunicamycin (5 μg/mL) at 37 °C for 30 min, harvested by centrifugation and washed with ice-cold ddH₂O. Total RNA extraction and cDNA synthesis by reverse transcription were performed as previously described [82]. To detect spliced and unspliced HAC1 mRNA, cDNA was used as a template in a PCR as follows: 95 °C for 5 min; 30 cycles consisting of 30 s at 95 °C, 30 s at 55 °C and 30 s at 72 °C; and 10 min at 72 °C [83]. The primers used are listed in Table S2 (in the Supplementary Materials). The PCR products of spliced and unspliced HAC1 mRNA were 189 and 208 bp, respectively, and separated by a 3% DNA agarose gel.

4.6. Real-Time Quantitative PCR (qPCR)

For real-time qPCR to detect UPR gene expression, the StepOne Plus™ real-time PCR system (Applied Biosystems, Framingham, MA, USA) was used. Each 15-μL reaction mixture contained 30 ng cDNA, 300 nM each forward and reverse primer, and 7.5 μL Power SYBR green PCR master mixture (Applied Biosystems). The primers used are listed in Table S2. The reactions were performed with 1 cycle at 95 °C for 10 min, followed by 40 repeated cycles at 95 °C for 15 s and 60 °C for 1 min. The PMA1 transcripts were used as an endogenous control for qPCR [83]. All experiments were repeated independently at least three times, and the average C_T values were determined. The relative fold change in the expression of each gene was calculated using the 2^–ΔΔCT method [84].

4.7. Measurement of Intracellular Reactive Oxygen Species (ROS)

To monitor intracellular ROS accumulation, the cell-permeant reagent H₂DCFDA was used as a probe for measuring ROS. H₂DCFDA is cleaved by intracellular esterase to become DCFH, which is later oxidized by ROS to form fluorescent DCF [85]. Briefly, cells treated with or without different concentrations of LL-37 were harvested by centrifugation and washed twice with sterile PBS. The cells were then incubated with 20 μg/mL H₂DCFDA at 30 °C for 30 min. The cells were washed three times with ice-cold PBS, and the fluorescence intensity was measured using a flow cytometer (BD Accuri^® C6, BD Biosciences) with the FL1 filter.

4.8. Lipid Peroxidation Assay

To determine the extent of lipid peroxidation, the levels of MDA were measured. C. albicans cells were treated with or without LL-37 (8 μg/mL) for 30 min. Subsequently, cells were harvested, washed twice with PBS and resuspended in PBS containing acid-washed glass beads. The cells were vortexed for 30 s and cooled on ice for 30 s, and this process was repeated 10 times. Cell debris was removed by centrifugation at 4 °C for 10 min, and the supernatant was collected. Next, lipid peroxidation was measured using the TBARS (TCA method) Assay Kit (Cayman Chemical, Ann Arbor, MI, USA) according to the manufacturer’s instructions. In this assay, TBARS formed by the reaction of MDA and thiobarbituric acid (TBA) under high temperature (90–100°C) and acidic conditions was measured colorimetrically at 530 nm.

4.9. Construction of the Ero1-Flag Expressing Strains

To construct the Ero1-Flag-expressing strains, the DNA sequence of Flag-tag was independently fused to one allele of the ERO1-coding sequence (at its 3′-end) in the wild-type (WT), sfp1Δ/sfp1Δ and SFP1-reintegrated strains. The strains generated were designated WT+Ero1-Flag, sfp1Δ/sfp1Δ+Ero1-Flag and SFP1-reintegrated+Ero1-Flag.

For strain construction, the SAT1-flipper method was used [77]. A KpnI-XhoI DNA fragment composed of the 3′-end of ERO1 was PCR-amplified from the SC5314 genome with the primer pair ERO1UR-F-KpnI and ERO1UR-R-Flag-XhoI (Table S2). Similarly, another SacII-SacI DNA fragment composed of the 3′ flanking region of ERO1 was amplified from the SC5314 genome with the primer pair ERO1DR-F-SacII and ERO1DR-R-SacI (Table S2). The resulting DNA fragments were independently cloned into the pSFS2A vector [77] to generate pSFS2A-ERO1-Flag. A large DNA fragment carrying the SAT1-flipper cassette flanked by the two PCR-amplified DNA fragments was then excised from pSFS2A-ERO1-Flag by KpnI/SacI digestion. The linear DNA fragment was purified and transformed into C. albicans cells for integration into the chromosome via homologous recombination. The transformants were selected for nourseothricin (Nou) resistance and verified by PCR. To pop out the SAT1-flipper cassette, cells were grown overnight at 30 °C in 5 mL of YPM medium to induce MAL2 promoter-regulated recombinase. Overnight-cultured cells were plated on YPM agar plates and incubated at 30 °C for 2 days. Single colonies were then screened for Nou resistance by streaking them on both YPD and YPDNou plates, and the Nou-sensitive strains were picked for PCR to verify the pop out of the SAT1-flipper cassette.

4.10. Detection of the Redox State of Ero1

To assess the redox state of Ero1, cells were treated with or without LL-37 (8 μg/mL) and tunicamycin (5 μg/mL) for 30 min, harvested by centrifugation, and washed twice with PBS. Protein extraction was performed as described above, and the protein concentration was measured using the Bradford Protein Assay (Bio–Rad, Hercules, CA, USA).

Proteins were mixed with 4x nonreducing sample buffer (250 mM Tris-HCl [pH 6.8], 40% glycerol, 8% SDS and 0.4% bromophenol blue) containing 25 mM 4-acetamido-4′-maleimidylstilbene-2,2′-disulfonic acid (AMS) (A485; Thermo Fisher Scientific, Waltham, MA, USA). The mixture was placed on ice for 15 min, incubated at 37 °C for 60 min, and boiled for 2 min [47,56,86]. The samples were resolved by nonreducing 6% SDS–PAGE and transferred to a PVDF membrane. The proteins were probed with an anti-Flag mouse monoclonal antibody (F3165, Sigma–Aldrich). HRP-conjugated goat anti-mouse IgG (GTX213111–01; GeneTex) was used as the secondary antibody. The ratio of oxidative Ero1 to reductive Ero1 was analyzed using ImageJ software [81].

4.11. Protein Secretion and BSA Degradation Assay

To determine protein secretion, the total content of extracellular proteins was assessed as previously described [47]. Cells were treated with or without LL-37 (8 μg/mL) and tunicamycin (5 μg/mL) for 30 min. Subsequently, the cells were centrifuged at 12,000× g for 10 min to collect the supernatants. Afterward, the supernatants were ultrafiltered with an Amicon Ultra4 Centrifugal Filter Unit (Millipore, Burlington, MA, USA) to concentrate the samples, and protein concentrations were determined using the Bradford Protein Assay (Bio–Rad).

The function of secreted protein was also evaluated by BSA degradation as previously described [64,65]. C. albicans cells grown overnight were harvested by centrifugation and washed twice with PBS. Cells were then subcultured in YCB-BSA-YE medium containing LL-37 (8 μg/mL) or tunicamycin (5 μg/mL) and incubated at 30 °C. After centrifugation at 12,000× g for 10 min, the supernatant protein samples were collected, separated by 12% SDS–PAGE and visualized with Coomassie blue staining.

We are grateful to Joachim Morschhäuser (Universität Würzburg, Germany) for generously providing the pSFS2A vector.