Transcriptional adaptation of C. albicans to IEC infection includes dynamic metabolic shifts and indicates scavenging of host zinc

To characterize the interaction of C. albicans with intestinal epithelial cells (IECs) from early fungal adhesion to initial invasion (up to 6 h) and later translocation and damage phases (12–24 h), we conducted dual-species RNA sequencing of C. albicans-infected IECs over a 24 h time course. Intestinal epithelial cells (C2BBe1) were seeded and differentiated for 12 d after reaching confluency to form a polarized cell layer [23]. Fungal and host RNA was isolated from C. albicans and IECs alone as well as from co-incubated samples. Samples were taken before infection (0 h), at 45 min (0.75 h), 3 h, 6 h, 12 h, and 24 h. C. albicans showed a clear time-dependent transcriptional response with distinct clusters for each time point (Fig 1A). C. albicans samples cultured with and without IECs showed similar transcriptional responses and clustered together, indicating the fungal response in this experimental model was largely independent of the presence of host cells (Fig 1A).

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Fig 1. C. albicans shows transcriptional changes specific to the presence of IECs during infection.

(A) Principal component analysis for WT (SC5314) C. albicans RNA sequencing data. Samples clustered together depending on the time point regardless of the presence of IECs (enterocytes). (B) GO terms significantly overrepresented among infection-specific C. albicans DEGs (*positive regulation of filamentous growth of a population of unicellular organisms in response to chemical stimulus; **biological process involved in interspecies interaction between organisms). Gene ratio represents the proportion of genes within each category that were significantly differentially expressed during infection compared to C. albicans cells only. The color scale represents the mean Log₂(fold-change) of significant DEGs within each category. Red indicates that most DEGs showed increased expression during infection, blue indicates most DEGs showed decreased expression, and grey indicates a mix of DEGs with increased and decreased expression within the category. Dots are only shown for significantly enriched categories (overrepresentation analysis, Benjamin-Hochberg adjusted p-value ≤ 0.05, S2 Table). (C) Gene expression of zinc-related genes comparing C. albicans cell during infection of IECs to those cultured in medium only. The color scale represents the Log₂(fold-change) for each gene during infection of IECs compared to medium only, with red representing higher expression during infection and blue representing lower expression during infection. Genes for the zinc transporters ZRT101, ZRT2, ZRT3 and ZRC1; the zinc-scavenging protein PRA1; and the transcription factor ZAP1 (CSR1) were less expressed during infection of IECs. An asterisk indicates the time points with a statistically significant difference in gene expression (DESeq2 p < 0.05). The mean MRN values are given in S1 Table.

https://doi.org/10.1371/journal.ppat.1012031.g001

To explore the transcriptional changes in C. albicans that are specific for the presence of IECs, we identified infection-specific differentially expressed genes (DEGs). For that, we compared fungal transcript levels in the presence vs absence of IECs at each respective time point (S1 Table). Using these DEGs, we performed functional enrichment analysis of gene ontology terms (GO; biological process category; S2 Table and Fig 1B) to identify the molecular patterns of infection.

We first looked at stress- or virulence-related functional categories that were enriched upon infection. In the earliest stages of infection, at 45 min, the enrichment analysis detected the term “response to chemical” which contained the stress-related catalase-encoding gene, CAT1, and the thioredoxin-encoding gene, TRR1 (S2 Table and Fig 1B). Decreased mRNA levels of these genes in infected samples compared to medium-only controls (S1 Table) suggests that C. albicans does not experience significant stress upon early contact to IECs. In late infection stages, however, the category “detoxification” was significantly overrepresented among infection-specific DEGs with increased transcript levels during infection (Fig 1B). These included many genes encoding predicted membrane transporters like the putative ABC transporter, CDR4, and the putative MFS transporter, QDR3 (S2 Table). This indicates increased stress related to the host or to host-cell damage.

The majority of enriched GO terms did not contain classical virulence-associated categories and did not include genes related to fungal filamentation or virulence. Only few enriched terms, including some related to lipid metabolism and cell adhesion, fall into these categories. Genes involved in lipid metabolism were significantly overrepresented at 6 h among the infection-specific DEGs (Fig 1B). These included many genes needed for ergosterol biosynthesis, which showed increased expression during infection of IECs (S1 and S2 Tables). As ergosterol biosynthesis is important for filamentation, this increased expression may indicate infection-specific surface changes in invading hyphae [24]. Enrichment of the category “cell adhesion” at 12 h similarly points to infection-specific surface changes (Fig 1B). This category includes DEGs coding for adhesins, cell surface proteins and cell wall integrity-related genes, such as EAP1, ALS1, PRA1, HWP1, XOG1, MNT2 and PMT1 (S1 and S2 Tables).

Most infection-specific changes were, however, due to metabolic adaptations of C. albicans. At time points of initial hypha formation prior to substantial host cell invasion (3 h) but also during host cell invasion and damage (12 h and 24 h) [11], the enrichment analysis revealed adaptations in the central fungal carbon metabolism in categories for carbohydrate transmembrane transport, the glyoxylate cycle, and monocarboxylic acid metabolism were enriched at 3 h (Fig 1B). These categories cover glucose transporter genes like HGT2 and HGT12, which are known to be repressed by high glucose levels [25], and genes involved in utilization of non-glucose carbon sources via the glyoxylate cycle and fatty acid beta oxidation, like ICL1, MLS1, FOX2, and PEX5 (S2 Table). Lower transcript levels of these genes during infection (S1 Table) indicates that the presence of IECs provides glucose to infecting C. albicans cells even before substantial fungal invasion. In addition, there was an infection-specific increase in translational activity at 3 h, with higher transcript levels of genes within the categories for ribonucleoprotein biogenesis and snRNA metabolic processes (Fig 1B). This indicates infection-specific re-organization of central cellular processes at an early time point.

At 12 h, genes falling under the term “pyruvate metabolic process” were also significantly enriched, including genes relevant for glycolysis (ENO1, CDC19, TDH3, and FBA2) and for the pyruvate dehydrogenase complex (PDA1, PDB1, PDX1, and LAT1) (Fig 1B and S2 Table). The lower transcript levels of these genes points to reduced glucose availability during these later stages of IEC infection (Fig 1B and S1 Table). Finally, at 24 h, the term “monosaccharide catabolism” was overrepresented among the infection-specific DEGs. This suggests that non-glucose C6 carbon sources are available specifically during the host-cell damage phase of infection (Fig 1B). These genes included some involved in galactose metabolism (GAL1, 7,10) and glycolysis (PFK1, 2) which were more highly expressed in the presence of IECs (S1 and S2 Tables). Translation-related processes were again enriched during infection at the 24 h time point (Fig 1B). Potentially, this reflects an adaptation to the more complex environment when C. albicans gains access to the host cell content.

Taken together, our data indicate that C. albicans dynamically adapts to the high availability of its preferred carbon source glucose at 3 h, to glucose limitation at 12 h, and the availability of alternative carbon sources upon host-cell damage at 24 h.

Importantly, not only did our analysis pick up infection-specific changes in macronutrient metabolism, but also adaptation to micronutrient availability. At 12 h, the term “zinc ion transport” was significantly overrepresented among the infection-specific DEGs (Fig 1B). This term contained the zinc transporter genes ZRT101, ZRT2, ZRT3, and ZRC1, which all had significantly reduced transcript levels during IEC infection. The zincophore gene PRA1 and the zinc homeostasis regulator gene ZAP1 (CSR1) showed the same pattern (Fig 1C) [26]. While mRNA levels of these zinc-related genes increased over time in both infecting C. albicans and C. albicans in medium only, their levels were higher in the absence of IECs, especially at later stages of infection (S1 Fig). The presence of their substrates strongly down-regulates the transcription of genes involved in micronutrient acquisition, like those for zinc [27–29]. Together, our data therefore indicate a stronger starvation for zinc when C. albicans is grown in the absence of IECs. In contrast, other metal-related terms were not overrepresented except for copper ion transport at initial infection stages (45 min) (Fig 1B). In fact, increasing transcript levels of iron-metabolic genes, like the high-affinity iron permease genes FTR1 and FTR2, in both infection and control conditions indicated a general iron starvation response at later time points. Unlike for zinc, this was not rescued by the presence of IECs (S1 Table).

Overall, our transcriptome analysis indicates that C. albicans adapts its metabolism and surface to the dynamic environmental changes during experimental IEC infection, including acquisition of zinc from the host cells.

Zinc acquisition during infection of IECs is ECE1-dependent

Zinc ion transport was a significantly overrepresented category among infection-specific DEGs during the later stages of invasion and host-cell damage (Fig 1B). This was not the case for genes involved in the transport of other metals or micronutrients, so we hypothesized that zinc may be of particular importance. To determine whether zinc acquisition plays a role during C. albicans infection of IECs, we utilized gene deletion mutants for the zinc importer genes ZRT101 and ZRT2, the zincophore gene PRA1, and the intracellular zinc transporter gene ZRC1. Zrt101, Zrt2, and Pra1 are involved in zinc acquisition and are upregulated by low zinc [27,28]; in contrast, Zrc1 is involved in zinc detoxification, although its transcriptional regulation has not been investigated. Loss of ZRT101, ZRT2, and PRA1 had no impact on adhesion to, invasion of, or translocation through IECs by zinc-pre-starved C. albicans cells (Fig 2A–2C). The strain lacking ZRC1, however, showed decreased invasion and strongly impaired translocation (Fig 2B and 2C). The dramatic decrease in translocation ability (~10-fold) is unlikely to be solely due to the decreased hypha formation (~1.5-fold) of this strain (S2 Fig). In contrast to these results, we found that loss of ZRT101, PRA1, and ZRC1 results in significantly decreased damage potential, and an almost complete absence of host-cell damage after deletion of both ZRT101 and ZRT2 (Fig 2D). The host-cell damage defect of this zinc uptake impaired mutant (zrt101ΔΔ/zrt2ΔΔ), but not of the zinc detoxification defective mutant (zrc1Δ/Δ) was rescued by addition of 25 μM ZnSO₄, a zinc concentration known to promote C. albicans growth while not being toxic (Fig 2D) [28]. This restoration of host-cell damage is likely due to increased growth and filamentation as assessed by microscopy images showing that without addition of zinc there is little-to-no fungal growth for the zrt101ΔΔ/zrt2ΔΔ strain (Fig 2E). Therefore, efficient zinc acquisition by C. albicans appears necessary to fully damage IECs, likely by fostering its growth and filamentation.

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Fig 2. Loss of zinc acquisition genes in C. albicans decreases damage potential by impairing growth during infection of IECs.

(A) Loss of zinc acquisition and storage genes had no effect on the adhesion of C. albicans to C2BBe1:HT29-MTX intestinal epithelial cells after 3 h. (B) Zinc acquisition and storage genes did not significantly affect the percentage of invasive hyphae after 6 h co-incubation with C2BBe1:HT29-MTX cells. The invasion of zrc1Δ/Δ was reduced, though not to a statistically significant degree. (C) Loss of ZRC1 significantly impaired translocation of C. albicans cells through C2BBe1:HT29-MTX intestinal epithelial cells grown on a porous membrane after 24 h. (D) The host-cell damage of the zrt101Δ/Δ, zrc1Δ/Δ, pra1Δ/Δ, and zrt101ΔΔ/zrt2ΔΔ strains was significantly reduced after 24 h of infection. The severe damage impairment could be rescued with exogenous addition of 25 μM ZnSO₄ during infection for zrt101ΔΔ/zrt2ΔΔ (P < 0.0001), but not for zrc1Δ/Δ (P > 0.9999). LDH release was adjusted by subtracting the release from uninfected and untreated host cells. (E) The addition of exogenous zinc during infection of C2BBe1:HT29-MTX cells increases the fungal growth of the zrt101ΔΔ/zrt2ΔΔ strain to a level similar to that of the WT (BWP17) strain. Fungal hyphae (blue) were stained with calclofluor white (scale bar = 100 μm). All values are shown as the mean with standard deviation. Invasion (B) and translocation (C) data were compared using a one-way analysis of variance (ANOVA) and the host-cell damage (D) data were compared using a two-way ANOVA. Statistical significance was determined with a post-hoc Dunnett’s multiple comparisons test. Statistical significance: *, P ≤ 0.05; ***, P ≤ 0.001; ****, P ≤ 0.0001.

https://doi.org/10.1371/journal.ppat.1012031.g002

We have shown that C. albicans experiences more intense late-stage zinc starvation in the absence of host cells–at time points at which damage of the IECs takes place in infected samples (Fig 1C) [11]. We therefore hypothesized that C. albicans acquires zinc from host cells during infection of the intestinal epithelium and that host-cell damage enables this zinc acquisition. The fungal peptide toxin candidalysin, encoded by the ECE1 gene, is the main damaging factor of C. albicans [11,14]. To further investigate the interplay between host-cell damage and zinc acquisition and the involvement of candidalysin in this process, we compared the transcriptional zinc starvation response of wild-type (WT, BWP17) C. albicans to an ece1Δ/Δ strain. The strain lacking ECE1 showed significantly higher transcript levels of ZRT2, PRA1, and ZRT3 after 24 h of infection of IECs, consistent with more severe zinc starvation (Fig 3B, 3C, 3D and 3F). A similar pattern was seen for ZRT101 mRNA levels, though the difference was not statistically significant (Fig 3A and 3F). When cultured in the absence of host cells, the WT (BWP17) and ece1Δ/Δ strains showed no significant differences in transcript levels of zinc-related genes (Fig 3F and S3A–S3E Fig). The addition of exogenous zinc during co-incubation with IECs did not affect the host-cell damage, fungal translocation, or fungal mass of the WT(BWP17) or ece1Δ/Δ strains during infection of IECs (S4A–S4C Fig). The differences in the transcriptional zinc starvation response between the WT (BWP17) and ece1Δ/Δ strains after 24 h were reduced with addition of zinc, though there was still a trend towards increased transcript levels in the ece1Δ/Δ strain (S4D Fig). These data indicate, that ECE1 is necessary for zinc acquisition during interaction with IECs.

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Fig 3. Loss of ECE1 increases the transcriptional zinc starvation response during infection of IECs.

Fold change in gene expression in C. albicans WT (BWP17) and ece1Δ/Δ strains during co-incubation with C2BBe1 cells for (A) ZRT101, (B) ZRT2, (C) PRA1, (D) ZRT3, and (E) ZRC1. Fold changes are calculated as the normalized gene expression at each time point compared to the respective 0 h control samples. (F) Fold change in normalized gene expression of the ece1Δ/Δ strain compared to WT(BWP17) at 24 h in medium only or during infection of IECs. Gene expression was analyzed by q-RT-PCR and is normalized to ACT1 as a housekeeping gene. All values are shown as the mean with standard deviation. All the ratio data from q-RT-PCR experiments were log-transformed before performing a two-tailed, paired t-test. Statistical significance: *, P ≤ 0.05.

https://doi.org/10.1371/journal.ppat.1012031.g003

IEC response to C. albicans infection by NFκB, MAPK, and TNF signaling and damage-dependent c-Fos/IL-8 induction

Similar to C. albicans, IECs showed a time-dependent transcriptional response with patterns reflecting the different stages of infection (Figs 1A and 5A). The transcriptomes of infected host cells differed from those cultured without C. albicans at all time points, showing a clear infection-specific response (Fig 4A). The time-dependent response of IECs presented as a drastic increase in the number of infection-specific DEGs after 12 and 24 h–time points at which C. albicans hyphae damage the epithelium and translocate (S1 Table) [11]. KEGG (S2 Table and Fig 4B) enrichment analysis was performed on the infection-specific IEC DEGs. Similar to C. albicans, there were few significant infection-specific pathways at the 45 min time point (overrepresentation analysis, adjusted p ≤ 0.05; Fig 4B). Components of the JNK and p38 MAPK pathway were induced upon early infection, including the c-Jun N-terminal kinase (JNK) gene MAPK10 as well as the transcription factor AP-1 components FOS, JUN and JUNB. FOS transcript levels remained high during infection at all later time points (S1 and S2 Tables).

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Fig 4. The transcriptional response of IECs to C. albicans infection is largely damage- and filamentation-independent.

(A) Principal component analysis for IEC RNA sequencing data. Samples from IECs (enterocytes) cultured in medium only cluster together across all time points and separately from those co-incubated with C. albicans. (B) KEGG categories significantly overrepresented among infection-specific DEGs in IECs. Gene ratio represents the proportion of genes within each category that were significantly differentially expressed during infection compared to IECs only. The color scale represents the mean Log₂(fold-change) of significant DEGs within each category. Red indicates that most DEGs showed increased expression during infection, blue indicates most DEGs showed decreased expression, and grey indicates a mix of DEGs with increased and decreased expression within the category. Dots are shown only for significantly enriched categories (overrepresentation analysis, Benjamin-Hochberg adjusted p-value ≤ 0.05, S2 Table). (C) Significantly differentially expressed genes between ece1Δ/Δ and efg1ΔΔ/cph1ΔΔ C. albicans strains compared to their respective WT strains during infection of IECs. The color scale represents the Log₂(fold-change) for each gene during infection in the mutant strain compared to the respective WT strain, with red representing higher expression in the mutant strain and blue representing lower expression in the mutant strain. An asterisk indicates the genes with a statistically significant difference (DESeq2 p < 0.05). The mean MRN values are given in S1 Table. (D) Release of IL-8 by C2BBe1 cells infected with C. albicans strains possessing varying damage potentials. The non-damaging efg1ΔΔ/cph1ΔΔ and ece1Δ/Δ strains induced less IL-8 release, though only significantly for the efg1ΔΔ/cph1ΔΔ strains. Conversely, the high-damaging cht2Δ/Δ strains induced significantly more IL-8 release. (E) NFκB activation as measured by the DNA-binding activity of the p65 transcription factor. Infection with all C. albicans strains increased the DNA-binding activity of p65 compared to the uninfected C2BBe1 cells, though not to a statistically significant degree for the efg1ΔΔ/cph1ΔΔ and ece1Δ/Δ strains. All values are shown as the mean with standard deviation. The IL-8 (D) and NFκB activation (E) data were compared using a one-way ANOVA with a post-hoc Šidák’s multiple comparisons test. Statistical significance: *, P ≤ 0.05; **, P ≤ 0.01; ***, P ≤ 0.001.

https://doi.org/10.1371/journal.ppat.1012031.g004

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Fig 5. NFκB activation limits ECE1-dependent damage and paracellular translocation.

(A) Inhibition of NFκB activation using the high affinity NFκB activation inhibitor quinazoline (QNZ) increases the damage potential of C. albicans wildtype, but not for the non-filamentous efg1ΔΔ/cph1ΔΔ strain or the non-damaging ece1Δ/Δ strain towards C2BBe1 cells after 24 h. LDH release was adjusted by subtracting the release from uninfected and untreated host cells. (B) Inhibition of NFκB activation increases fungal translocation of C. albicans after 24 h across intestinal epithelial cells independent of damage potential. (C) Blocking NFκB activation significantly increases the breakdown of barrier integrity after 24 h of the intestinal epithelial cell layer for efg1ΔΔ/cph1ΔΔ and ece1Δ/Δ, with a similar but not significant effect on both WT strains. (D) E-cadherin protein levels normalized to GAPDH and presented relative to levels in untreated C2BBe1 cells. QNZ treatment further increased degradation of E-cadherin during infection with WT (SC5314) and ece1Δ/Δ. (E) Fluorescent labeling of E-cadherin during C. albicans infection with and without QNZ treatment (scale bar = 50 μm). Inhibition of NFκB activation decreased the organization of E-cadherin at IEC borders upon infection with the ece1Δ/Δ and both WT strains. Representative pictures for each strain and treatment condition are shown from 3 biological replicates with median score values in the inset for each strain and condition. All values are shown as the mean with standard deviation. Host-cell damage (A), fungal translocation (B), and barrier integrity (C) data were compared using a one-way ANOVA. Statistical significance for host-cell damage (A) and fungal translocation (B) data was determined with a post-hoc Tukey’s multiple comparisons test, while significance for the barrier integrity (C) data was determined using a post-hoc Šidák’s multiple comparisons test. Statistical significance: **, P ≤ 0.01; ****, P ≤ 0.0001.

https://doi.org/10.1371/journal.ppat.1012031.g005

Soon after initial hypha formation at 3 h, we observed an infection-specific increase in mRNA levels of epithelial genes involved in oxidative phosphorylation (Fig 4B). Specifically, transcript levels were higher for mitochondrial genes (ND1, 2, 4, 5, 6; ND4L; COX1, 2, 3; ATP6, 8; CYB), which has previously been observed in vaginal epithelial cells [19] (S1 and S2 Tables). After 6 h when invasion of host cells occurs, and until the final time point at 24 h, genes involved in innate immune responses were significantly overrepresented in the infection-specific DEGs. MAPK, TNF, NFκB, and IL-17 signaling pathways were overrepresented due to genes with increasing transcript levels during IEC infection (Fig 4B). Genes involved in the classical MAP kinase pathway (including growth factor genes EFNA1, EREG, AREG, TGFA; calcium voltage-gated channel CACNA genes; receptor tyrosine kinase genes NGFR, EPHA2, FGRF1; protein kinase C gene PRKCB; protein tyrosine phosphatase and mitogen-activated protein kinase phosphatase genes PTPN7, DUSP2, 4, 5, 6, 10; NFKB2) had increased transcript levels upon C. albicans infection (S1 and S2 Tables). Genes encoding chemokines (CXCL1, 2, 3, 8, 17), inflammatory cytokines (IL1B, IL11, IL12A, IL27, IL32 CCL20), as well as genes for intracellular signaling proteins (GADD45A, B; FOS; FOSL1; FOSB; JUN; A20) showed increased transcript levels upon C. albicans infection (S1 and S2 Tables). The late-stage induction of innate immune signaling pathways is consistent with data previously published for IECs [18].

The majority of infection-specific host DEGs only appear after fungal invasion and resulting host-cell damage. In comparison, vaginal epithelial cells show a uniform early response to various fungal species, but the response diverges at the later time points and is connected to damage potential [19]. We sought to determine whether the IEC DEGs at later time points reflect a general response to fungi or rather a specific response to C. albicans filamentation and damage. To determine which of the host responses identified were filamentation- or damage-specific, we extended our transcriptome analysis to compare IECs infected with C. albicans WT, non-damaging (ece1Δ/Δ), and non-filamentous and non-damaging (efg1ΔΔ/cph1ΔΔ) strains. The two mutant strains used (ece1Δ/Δ and efg1ΔΔ/cph1ΔΔ) were constructed in different genetic backgrounds, therefore the gene expression data was always compared to the corresponding WT strains (BWP17 and SC5314, respectively) to minimize the effect of strain-specific, genetic differences on the host response [30]. The set of genes that were only differentially expressed in response to the efg1ΔΔ/cph1ΔΔ strain was then categorized as a filamentation-specific response. Genes that were differentially expressed in response to both the efg1ΔΔ/cph1ΔΔ and ece1Δ/Δ strains, were considered to constitute a damage-specific response. The filamentation-specific response of IECs to C. albicans did not consist of many genes, though it included mitochondria-associated genes (COX7B, ND2, CYTB, ND4L) and some encoding heat-shock proteins (HSP90AA1 and HSP90AA6P), all with lower transcript levels in the non-filamentous mutant as compared to the WT (SC5314) (Fig 4C and S5 Fig).

The damage-specific response of IECs comprises even fewer genes, which in most cases showed lower transcript levels in IECs infected with the ece1Δ/Δ and efg1ΔΔ/cph1ΔΔ mutants than with the respective WT strains. According to these data, IECs respond to C. albicans-mediated damage via upregulation of EGR1 and EGR2, the FOS genes FOSL1 and FOSB as well as the IL-8-encoding gene CXCL8 (Fig 4C and S5 Fig). c-Fos is a major component of the oral epithelial cell (OEC) response to C. albicans filamentation and damage [13, 14]. Similarly, EGR1 induction as well as IL-8 secretion are known responses to C. albicans infection in OECs [31].

To test how conserved the response of IECs and OECs to C. albicans and fungus-mediated damage are, we performed western blotting for the OEC response components EphA2 and EGFR (receptors), the transcription factor c-Fos, the phosphatase MKP1 and the kinases Akt and p38 [13,16,17,32]. We observed an infection-specific increase in phosphorylation of EphA2 and Akt, as well as an infection-specific increase in c-Fos protein levels. c-Fos signals were consistently reduced in the ece1Δ/Δ mutant. While there was also decreased phosphorylation of EphA2 for the ece1Δ/Δ mutant compared to the WT, this was not consistent for all replicates (S6A and S6B Fig). Thus, we see a partial conservation in the damage response during C. albicans infection of IECs compared to OECs, with ECE1-dependent induction of c-Fos and potentially phosphorylation of EphA2, but no EGFR or MKP1 phosphorylation and no change in the phosphorylation of p38 or Akt in the absence of ECE1 compared to the WT [14–16,33,34].

To verify that production of IL-8 in IECs is filamentation- and damage-dependent, we measured IL-8 secretion in response to C. albicans infections. We compared the WT with the non-filamenting efg1ΔΔ/cph1ΔΔ strain, the non-damaging ece1Δ/Δ strain, and a previously identified strain (cht2Δ/Δ) with increased damage of IECs [11] (Fig 4D). Indeed, the efg1ΔΔ/cph1ΔΔ and ece1Δ/Δ mutants elicited less IL-8 secretion while cht2Δ/Δ induced more IL-8 secretion than their respective WT strains. Overall these data show that while the response of IECs to C. albicans infection is largely independent of hypha formation and fungal-mediated damage, there is a damage-specific aspect via c-Fos induction and IL-8 secretion.

Intestinal epithelial cells limit fungal damage and translocation via NFκB activation

Most of the pathways overrepresented during infection with WT C. albicans were also found during infection with the non-damaging and non-filamentous strains (S2 Table), again suggesting that the IEC transcriptional response to C. albicans is largely independent of fungal filamentation or damage. Among other signaling pathways that were triggered by the initiation of fungal-mediated damage (Fig 4B), DEGs were enriched in NFκB signaling after infection with all four C. albicans strains (S2 Table). Previous research has shown that C. albicans-induced NFκB activation in IECs limits the damage potential during infection [18]. We confirmed the C. albicans-induced NFκB activation in IECs and additionally showed that infection with C. albicans strains with high or low damage potential and even with a non-filamentous mutant elicit similar activation of NFκB by IECs (Fig 4E). This suggests that NFκB activation is a general epithelial response to the presence of C. albicans (Fig 4E).

To determine whether the previously described NFκB-dependent limitation of IEC damage is predominantly host- or fungal-driven, IECs were infected with the non-filamentous efg1ΔΔ/cph1ΔΔ and non-damaging ece1Δ/Δ strains in the presence and absence of the potent NFκB activation inhibitor quinazoline (QNZ) [18, 35]. In agreement with previous data, both tested WT strains elicited significantly more host-cell damage when NFκB activation was blocked (Fig 5A) [18]. In contrast, there was no statistically significant increase in damage caused by the strains lacking either ECE1 or both EFG1 and CPH1 (Fig 5A). Treatment of IECs with a DMSO vehicle control had no significant effect on either the damage potential or fungal translocation for the WT (BWP17) and ece1Δ/Δ strains (S7A and S7B Fig).

As host-cell damage of IECs is tightly linked to fungal translocation, the translocation rates of C. albicans in the presence of QNZ were also measured [11]. As expected from our host-cell damage data, blocking NFκB activation significantly increased the number of translocated fungal CFUs for both WT strains compared to the untreated controls (Fig 5B). The non-filamentous efg1ΔΔ/cph1ΔΔ strain showed no increase in its translocation rate, which was expected given that this strain cannot form hyphae or damage the host cells (Fig 5A and 5B) [11,36]. Surprisingly, despite showing no significant increase in host-cell damage upon QNZ-treatment [<50% of untreated WT (BWP17)], the ece1Δ/Δ strain showed a significant increase in translocation comparable to that of the WT (BWP17) under normal infection conditions (Fig 5B). This increased translocation in the absence of increased host-cell damage was confirmed for the ece1Δ/Δ strain with the use of another NFκB inhibitor with a different mode of action that directly inhibits the DNA-binding activity of the p65 subunit [37,38] (S8A and S8B Fig). These data together indicate that NFκB can limit C. albicans translocation, at least in part, independently of host-cell damage.

We have previously shown that C. albicans translocation across IECs occurs mainly via a transcellular route and is associated with filamentation- and candidalysin-dependent necrotic host-cell damage [11]. The increased translocation of the ece1Δ/Δ strain without any increased host-cell damage upon NFκB inhibition suggests that NFκB activation likely limits a non-damaging route for translocation (Fig 5B). To test this hypothesis, the transepithelial electrical resistance (TEER) of IECs was measured with and without QNZ treatment as a metric for barrier integrity. After 24 h, IECs infected with both WT strains showed decreased barrier integrity upon QNZ treatment, though not to a statistically significant degree (Fig 5C) [WT (SC5314), P = 0.2466; WT (BWP17), P = 0.0824]. These results match the increased host-cell damage and translocation data for both strains. There was also a significant decrease in the barrier integrity for IECs infected with both, the efg1ΔΔ/cph1ΔΔ and ece1Δ/Δ strains upon QNZ treatment, again pointing to a host damage-independent effect of NFκB activation on translocation (Fig 5C). This was also confirmed for the WT and ece1Δ/Δ strains using the second NFκB inhibitor (S8A–S8C Fig).

Non-damaging routes of fungal translocation that still reduce barrier integrity can involve induction of host-cell apoptosis or the use of paracellular routes with degradation of cell-cell connections such as tight junctions or adherens junctions [11]. As the NFκB signaling pathway can also induce the expression of anti-apoptotic genes and we observed an overrepresentation of infection-specific DEGs for IECs involved in apoptosis at 12 h when fungal-mediated damage begins (Fig 4B) [39]. We therefore tested whether blocking NFκB activation during infection with C. albicans increases apoptosis or other types of programmed cell death in IECs. IECs were left untreated or treated with QNZ, a pan-caspase inhibitor (Z-VAD), or both, and were then infected with different C. albicans strains. Treatment with Z-VAD alone had no significant effects on host cells damage, loss of barrier integrity, or fungal translocation (S9A–S9C Fig). Similarly for IECs with blocked NFκB activation, addition of Z-VAD led to no significant differences with any of the strains tested (S8A–S8C Fig). Together, these data suggest that the ability of IECs to limit cellular damage and translocation through NFκB activation is dependent on the filamentation and damage potential of C. albicans itself, and does not rely on increased apoptosis or other forms of programmed cell death.

To finally test whether the decreased barrier integrity during infection with the ece1Δ/Δ strain was associated with increased degradation of cell-cell junction proteins consistent with increased paracellular translocation, epithelial barriers were stained for E-cadherin 24 h after infection with and without QNZ treatment. Micrographs from three biological replicates were then scored by a blinded observer for the consistency of the cell-cell borders and overall organization of the E-cadherin staining. Non-inhibitor treated uninfected and efg1ΔΔ/cph1ΔΔ-infected IECs showed a uniform staining of E-cadherin, indicating an intact host cell layer (Fig 5C and 5E). In contrast, untreated cells infected with either WT strain or the ece1Δ/Δ strain showed a weaker staining (Fig 5E). Inhibitor treatment further decreased the E-cadherin signal in IECs infected with either of the two WT strains, as expected from the inhibitor-dependent increase in damage (Fig 5A). Importantly, the ece1Δ/Δ strain showed less intact host cells and decreased fluorescence upon NFκB activation inhibition, while the host cell layer was largely unchanged for the uninfected and efg1ΔΔ/cph1ΔΔ-infected IECs when treated with QNZ (Fig 5E). These data indicate that NFκB limits paracellular translocation by stabilizing cell-cell junction proteins like E-cadherin. They also suggest that, upon NFκB inhibition, the physical presence of hyphae can break down intercellular barrier function even in the absence of toxin-mediated damage.

To further investigate the effects of NFκB inhibition on cell-cell junction proteins at a molecular level, we measured the junction proteins E-cadherin and claudin-1 by western blotting for infected IECs with and without QNZ treatment (S10A Fig). Western blotting showed that the total protein levels of E-cadherin and claudin-1 (normalized to GAPDH protein levels) in C. albicans infected IECs were largely unaffected by NFκB inhibition (Figs 5D and S10B Fig). There was no consistent difference in E-cadherin levels upon inhibitor treatment during infection with the WT (BWP17) or efg1ΔΔ/cph1ΔΔ strains. However, relative E-cadherin levels upon treatment were reduced for IECs infected with the WT (SC5314) and ece1Δ/Δ strains, though not to a statistically significant degree (Fig 5D). The same trend was true for the ece1Δ/Δ and efg1ΔΔ/cph1ΔΔ strains for claudin-1 (S10B Fig). Though the decrease in both E-cadherin and claudin-1 for the ece1Δ/Δ strain was not significant, it did correlate with the decrease in barrier integrity (Fig 5C) and is also consistent with the organization of the cell-cell junction proteins (Fig 5E). In summary, these data show that NFκB activation limits ECE1-mediated host-cell damage and independently limits fungal translocation. It does do by maintaining cell-cell junctions and the epithelial barrier integrity in the absence of direct fungal-mediated host-cell damage.

Candida albicans strains and growth conditions

C. albicans strains used in this study are shown in Table 1. The WT strains SC5314 and BWP17+CIp30 are referred to as WT (SC5314) and WT (BWP17), respectively. Strains were routinely cultivated on/in YPD agar/broth (1% yeast extract, 2% peptone, 2% D-glucose with or without 1.5% agar) at 30°C. Overnight (O/N) cultures were cultured for 16 h in YPD broth at 30°C with shaking at 180 rpm unless otherwise specified. Cells were then washed twice with phosphate-buffered saline (PBS) and the cell number was adjusted.

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Table 1. C. albicans strains used in this study.

https://doi.org/10.1371/journal.ppat.1012031.t001

For experiments with prior zinc starvation, C. albicans strains were grown for 24 h in YPD liquid medium supplemented with 1 mM ZnSO₄ at 30°C with shaking at 180 rpm. The OD₆₀₀ was then adjusted to 0.5 in SD liquid medium supplemented with 1 mM ZnSO₄ and incubated at 30°C for 24 h with shaking at 180 rpm. The cells were then washed twice with 1 mM EDTA, washed twice with water, and then diluted to an OD₆₀₀ of 0.5 in Zn-free SD liquid medium in acid-washed, plastic flasks. The fungal strains were then incubated for another 24 h at 30°C with shaking at 180 rpm.

Culture of intestinal cells

The intestinal epithelial Caco-2 brush border expressing 1 cell line (C2BBe1; ATCC, CRL2102) [68] and the human intestinal goblet cell line (HT29-MTX; ATCC, HTB-38; CLS, Lot No. 13B021) were routinely cultivated in Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco, Thermo Fisher Scientific) supplemented with 10% fetal bovine serum (FBS) (Bio&Sell), 10μg/ml Holotransferrin (Calbiochem, Merck), and 1% non-essential amino acids (Gibco, Thermo Fisher Scientific) at 37°C with 5% CO₂ for no longer than 15 passages. C2BBe1 cells were seeded in 6-well plates at a concentration of 5×10⁵ cells/well for RNA isolation. C2BBe1 and HT29-MTX cells were seeded in 96-well plates and transwell inserts (polycarbonate membrane with 5 μm pores; Corning) at a 70:30 ratio (C2BBe1:HT29-MTX) and a total concentration of 2×10⁴ cells/well or insert for damage and translocation assays, respectively. For adhesion and invasion assays, C2BBe1 and HT29-MTX cells were seeded in 24-well plates with coverslips at a 70:30 ratio (C2BBe1:HT29-MTX) at a total concentration of 1×10⁵ cells/well. All well plates and transwell inserts were coated with collagen I (10 μg/ml for 2 h at room temperature; Thermo Fisher Scientific) and maintained for 12 d at 100% confluency for differentiation with regular medium exchange before infection. Just prior to infection with C. albicans, the medium was removed and fresh DMEM without FBS, Holotransferrin, or 1% non-essential amino acids was added to the cells.

RNA isolation

C2BBe1 cells were cultured for 12 d in collagen-coated 6-well plates. To prepare samples for dual-RNA sequencing and qPCR, the differentiated intestinal epithelial cells (IECs) were infected with 2×10⁶ Candida cells/well and incubated at 37°C and 5% CO₂ for 0.75, 3, 6, 12, or 24 h in serum-free DMEM medium. For qPCR samples with added zinc, the fungal and C22Be1 cells were co-incubated in serum-free DMEM with 25 μM ZnSO₄ for 24 h. In parallel, C2BBe1 and C. albicans cells were cultured individually in 6-well plates in serum-free DMEM medium. C. albicans cells from the O/N cultures were collected as uninfected controls for the fungus (0 h C. albicans), and uninfected IECs were harvested as well (0 h IECs). At each respective time point, the medium was removed, unadhered Candida cells were washed away, and 650 μl of RLT buffer (RNeasy Minikit; Qiagen) was added to each well. The plates were frozen with liquid nitrogen and thawed at room temperature (RT). Each well was scraped and the suspensions were centrifuged for 8 min (20,000×g) to pellet the Candida cells. The supernatant was removed and the host RNA was isolated using the RNeasy Minikit (Qiagen) according to the manufacturer’s instructions. The fungal RNA was isolated from the pellet using a freeze-thaw method described previously [69]. RNA concentrations were measured using a NanoDrop 1000 Spectrophotometer (Thermo Fisher Scientific) and the quality was controlled using a 2100 Bioanalyzer (Agilent Technologies).

Fungal RNA used for q-RT-PCR was isolated as described previously with minor modifications [69]. Briefly, fungal pellets were resuspended in AE buffer (50 mM Na-acetate pH 5.3, 10 mM EDTA) with 0.5% SDS and transferred to screw cap tubes with acid-washed glass beads. An equal volume of phenol/chloroform/isoamylalcohol (25:24:1) was added and the fungal cells were lysed using a FastPrep-24 5GMP Biomedicals. AE buffer was added and the samples were centrifuged for 10 min (20,000×g, 4°C). The aqueous phase was transferred to a PLG-heavy tube (QuantaBio), re-extracted with an equal volume of phenol/chloroform/isoamylalcohol, and centrifuged for 5 min (20,000×g, 4°C). The RNA was then precipitated overnight at -20°C with pure ethanol and sodium acetate. The RNA concentration was measured using a NanoDrop 1000 Spectrophotometer (Thermo Fisher Scientific) and samples were stored at -70°C.

Fungal gDNA isolation

Differentiated C2BBe1:HT29-MTX cells in 24-well plates were infected with 4×10⁵ C. albicans cells and incubated at 37°C with 5% CO₂ for 24 h in serum-free DMEM with or without 25 μM ZnSO₄. The medium was removed, the host cells were lysed with 350 μl RLT buffer (RNeasy Minikit: Qiagen), and the fungal cells were spun down. Fungal gDNA was isolated as previously published with the following modifications [70]. The fungal pellet was washed with water once. Equal volumes of lysis buffer (2% Triton X-100; 1% SDS; 100 mM NaCl; 10 mM Tris, pH 8; 1 mM EDTA) and phenol:chloroform:isoamylalcohol (25:24:1) were added and the samples were transferred to tubes with acid-washed glass beads. The fungal cells were disrupted using a FastPrep-24 5G (MP Biomedicals). Half the total volume of TE buffer (10 mM Tris; 1 mM EDTA; pH 7.5) was added. Nucleic acid was precipitated from the aqueous phase and resuspended in 400 μl TE buffer following RNase A treatment (10 mg/ml; Sigma) for 30 min at 37°C. gDNA was precipitated in pure ethanol, washed and resuspended in 25 μl nuclease-free water.

cDNA synthesis and q-RT-PCR

Isolated RNA (1μg) was treated with DNase I (Baseline-ZERO DNase, Lucigen) according to the manufacturer’s instructions and then transcribed into complementary DNA using 50μM of oligo (dT)_12-18 primer (Invitrogen), 150 U of Superscript III Reverse Transcriptase (Invitrogen), and 10 U RNase OUT (Invitrogen). The cDNA was then diluted 1:5 and used for q-RT-PCR with the GoTaq qPCR Master Mix (Promega) in a CFX96 thermocycler (BioRad). The expression levels were normalized against the C. albicans ACT1 gene.

For fungal gDNA samples, EFB1 was amplified from 1μl of template with qPCR Master Mix (Promega) in a CFX Opus Real-Time PCR System (BioRad). All primers used are listed in Table 2. The relative gDNA for each sample was calculated relative to the untreated WT(BWP17) for the respective biological replicate.

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Table 2. Primers used for q-RT-PCR.

https://doi.org/10.1371/journal.ppat.1012031.t002

RNA sequencing and transcriptional analysis

Individual C. albicans and human C2BBe1 samples were combined for dual-RNA sequencing for uninfected isolated samples for time course data. Both C. albicans and Homo sapiens reads were present in infected samples. C2BBe1 reads were present in infection data for mutant vs. WT comparison.

Library preparation and RNA sequencing were carried out at Eurofins Genomics GmbH (Ebersberg, Germany) using the Illumina HiSeq 2500 platform (time course analysis) or Illumina HiSeq 4000 platform (C. albicans mutant to WT comparisons). For library preparation, after poly(A) enrichment, mRNA was fragmented and random-primed cDNA synthesis was performed followed by adaptor ligation and adaptor-specific PCR amplification. Single sequence reads of 50 bp were produced.

Preprocessing of raw reads including quality control and gene abundance estimation was done with the GEO2RNaseq pipeline (v0.100.3) [71] in R (version 3.6.3). Quality analysis was done with FastQC (v0.11.5) before and after trimming. Read-quality trimming was done with Trimmomatic (v0.36). Reads were rRNA-filtered using SortMeRNA (v2.1) with a single rRNA database combining all rRNA databases shipped with SortMeRNA. Reference annotation was created by extracting and combining exon features from corresponding annotation files. Reads were mapped against the joined reference genomes of H. sapiens (Ensembl_GRCh38) and C. albicans (C_albicans_SC5314_version_A22) using HiSat2 (v2.1.0, single end mode). Gene abundance estimation was done with featureCounts (v2.0.1) in single-end mode with default parameters. MultiQC version 1.7 was finally used to summarize and assess the quality of the output of FastQC, Trimmomatic, HiSat, featureCounts and SAMtools. The count matrices with gene abundance data without and with median-of-ratios normalization (MRN) were extracted. Raw files are accessible under the Gene Expression Omnibus accession number GSE237496 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE237496) [72].

Differential gene expression was analyzed using GEO2RNaseq. Pairwise tests were performed using four statistical tools (DESeq2 v1.26.0) to report p values and multiple testing corrected p values using the false-discovery rate method q = FDR(p). Mean MRN values were computed per test per group including corresponding log₂ of fold-changes. Gene expression differences were considered significant if they were reported significant by DESeq2 (q ≤ 0.05) and |log₂(fold-change[MRN based])| ≥ 1.

Functional enrichment analysis was performed using overrepresentation analysis. Gene ontology (Biological process) for C. albicans was performed by parsing data gene lists to the web application of FungiDB for gene annotation and subsequent analysis using default parameters (https://fungidb.org) [73]. Enrichment for human reads against KEGG pathways were computed using g:Profiler from within R (package gprofiler2, v0.2.1). Enriched terms were discarded if background size was below 3 or above 1000 genes. A bash script was used to call the Revigo web app for removing redundant terms (reduction factor = 0.5, removal of obsolete terms = yes) [74]. KEGG terms were further filtered to exclude coincidental significant hits that were not relevant for the current study (virus-related categories, “Organismal systems” except “Immune system”, “Human diseases”, “Drug development”). Programming code and data necessary to generate plots shown in this manuscript were deposited at Github: https://github.com/SchSascha/Cal_Translocation.

Quantification of adhesion, invasion, and hyphal length

Differentiated C2BBe1:HT29-MTX cells in 24-well plates with glass coverslips were infected with 1×10⁵ C. albicans cells and incubated at 37°C with 5% CO₂ for 1 h (adhesion) or 6 h (invasion). For adhesion, samples were fixed with 4% formaldehyde and washed twice with PBS. Adherent C. albicans cells were stained with 10 μg/ml Calcofluor white (CFW) (Fluorescent Brightener, Sigma-Aldrich) in 100 mM TRIS-HCl (pH 9.5) for 30 min at RT. After washing with water, samples were mounted on a glass slide with mounting medium (ProLong Gold Antifade, Invitrogen) and imaged. For each replicate, 12 representative images were counted and the mean number of adherent cells in a defined area was calculated from four independent replicates. For quantification of invasion, extracellular C. albicans hyphae were stained with CFW as stated above. The host cells were then permeabilized with 0.5% Triton X-100 for 5 min and washed with PBS. Intracellular C. albicans hyphae were first labelled with 20–25 μg/ml rabbit anti-C. albicans antibody (Acris) for 3 h at 4°C. After washing with PBS, 4 μg/ml goat anti-rabbit antibody conjugated to AlexaFluor 488 was added and incubated at 37°C for 1 h. The samples were washed with PBS and mounted as mentioned above. The number of invasive hyphae was determined from 100 hyphae counted per sample. All samples were imaged using an Axio Observer fluorescence microscope (Zeiss). To quantify hyphal length, 2×10⁴ C. albicans cells were seeded per well in a 96-well plate in DMEM and incubated for 3 h at 37°C with 5% CO₂. Images were taken using a Cell Discoverer 7 microscope (Zeiss) and the hyphal length was measured using Zeiss Zen3.4 (blue edition).

Quantification of host cytotoxicity (LDH assay)

Differentiated C2BBe1:HT29-MTX cells in 96-well plates were infected with 8×10⁴ C. albicans cells and incubated at 37°C with 5% CO₂ for 24 h. Epithelial damage was quantified by release of lactate dehydrogenase (LDH) from IECs. LDH concentrations were measured using a cytotoxicity detection kit (Roche) according to the manufacturer’s instructions and LDH from rabbit muscle (Roche) was used to generate a standard curve. The baseline LDH concentration of uninfected and untreated IECs was subtracted and the corrected LDH release is shown as a concentration (ng/ml) present in the supernatant unless otherwise stated.

In vitro translocation assay

Differentiated C2BBe1 cells or a mix of C2BBe1:HT29-MTX cells in transwell inserts were infected with 1×10⁵ C. albicans cells and incubated at 37°C with 5% CO₂. Supernatant from the upper compartment was removed 24 hpi and used for LDH measurement as described above. The lower compartment was treated with 20 U/ml zymolyase (Amsbio) for 2 h at 37°C and 5% CO₂ to detach translocated hyphae. The zymolyase-treated hyphae were then plated on YPD agar, incubated at 30°C for 2 d, and the colony forming units (CFUs) were counted.

For experiments with NFκB inhibition, C2BBe1 cells were treated with DMEM with 2.5 μM 6-Amino-4-(4-phenoxyphenylethylamino)quinazoline (QNZ) (EVP4593; Sigma-Aldrich) or either 2.5 or 5 μM N-(6-benzoyl-1H-benzo[d]imidazol-2-yl)-2-(1-(thieno[3,2-d]pyrimidin-4-yl)piperidin-4-yl)thiazole-4-carboxamide (SC75741) (MedChemExpress) in the lower compartment for 1 h prior to infection at 37°C with 5% CO₂. IECs were then infected as above. After 24 h the trans-epithelial electrical resistance (TEER) was measured using a volt-ohm meter (EVOM2, World Precision Instruments), supernatants from the upper compartment were collected for LDH measurements, the lower compartment was treated as above to measure CFUs, and the transwell membranes were removed for staining. IECs were fixed with Histofix (Roth) for 10 min at 37°C then washed with PBS. The cells were permeabilized with 0.5% TritonX-100 for 10 min at RT and washed with PBS. Cells were blocked with 2% BSA for 10 min at 37°C, washed with PBS, and incubated with a primary mouse anti-E-cadherin antibody (10 μg/ml) (BD Biosciences) for 4 h at 4°C, then washed with 2% BSA. Cells were then stained with a secondary goat anti-mouse antibody conjugated to AlexaFluor 488 (10 μg/ml) (Invitrogen). Samples were washed with PBS, mounted on glass slides, and images with an Axio Observer fluorescence microscope (Zeiss). Samples were prepared from 3 independent biological replicates and 3 images were taken per strain per condition for each replicate. Images were randomized and scored by a blinded observer. Scores ranged from 1 to 5, with a score of 1 representing a sample with consistent, organized staining of E-cadherin at cell-cell borders throughout the sample. A score of 5 was assigned to samples with no visible staining of E-cadherin at cell-cell junctions. For experiments using the pan-caspase inhibitor (Z-VAD), host cells in transwell inserts were treated for 1 h prior to infection with 25 μM of the inhibitor alone or in combination with 2.5 μM of QNZ as described above.

For experiments with added zinc, the number of fungal cells was adjusted in either DMEM or DMEM + 25 μM ZnSO₄. Differentiated C2BBe1:HT29-MTX cells in transwell inserts were infected as above, with either DMEM or DMEM + 25 μM ZnSO₄ in the transwell insert and DMEM in the lower compartment. After 24 h incubation at 37°C with 5% CO₂, the translocated fungal CFUs were plated as described above.

Immunoblotting

For detection of junction proteins with and without NFκB inhibition, C2BBe1 cells were infected as mentioned for the in vitro translocation assay with and without the NFκB inhibitor QNZ and incubated for 24 h at 37°C with 5% CO₂. The transwell membranes were then removed, the host cells were scraped off, and spun down for 5 min at 4°C at 500×g. The pellets were lysed in RIPA lysis buffer (VWR) supplemented with a protease inhibitor mix (Roche) and Benzonase (Millipore) and chilled for 15 min on ice. The samples were spun down for 15 min at 4°C at 18,000×g. The supernatants were collected and stored at -70°C. The protein concentrations of total protein extracts were measured by a Lowry protein assay kit (Bio-Rad Laboratories GmbH). 25 μg of each sample was used for a standard 12% SDS polyacrylamide gel electrophoresis (SDS-PAGE) at 30 mA and 200 V. Therefore, lysates were supplemented with SDS sample buffer (50 mM Tris/HCl, 2% (v/v) glycerol, 2% (v/v) β-mercaptoethanol, 1.6% (w/v) SDS, 0.004% (w/v) Serva Blue G-250, pH 6.8) and boiled for 10 min at 95°C. After SDS-PAGE with subsequent blotting of the samples onto an Amersham Protran 0.45 μm nitrocellulose membrane at 1.8 mA/cm² and 25 V, blots were incubated with 5% (w/v) milk dissolved in TBS (50 mM Tris/HCl, 140 mM NaCl, pH 7.2) for 1 h at room temperature. For specific, immunologic labeling, primary antibodies against E-Cadherin (1:1000, MAB1838, R&D Systems), Claudin-1 D5H1D (1:1000, 13255, Cell Signaling Technology), and GAPDH D16H11 (1:1000, 5174, Cell Signaling Technology) were used in 5% (w/v) milk/TBS for 3 h at room temperature or 16–48 h at 4°C. After 3 times washing in TBS-T (TBS supplemented with 0.05% (v/v) Tween-20), membranes were incubated for 1.5–3 h with IRDye 800CW donkey anti-mouse or donkey anti-rabbit (1:10000, 925–32212 and 925–32213, LI-COR GmbH) as secondary antibodies, respectively. Immunological detections were carried out using an Odyssey M imaging system (LI-COR GmbH). Blots were analyzed by Image Studio Lite Version 5.0 (LI-COR GmbH).

For detection of immune signaling proteins, C2BBe1 cells were seeded in 6-well plates and infected with WT (BWP17) and ece1Δ/Δ C. albicans as described above for RNA isolation experiments. After 6 h of infection, tissue culture plates were placed on ice, the medium was removed, and the cells were washed with ice-cold PBS. Cells were lysed with 120 μL of RIPA buffer (25 mM Tris-HCl pH 7.4, 150 mM NaCl, 1% Nonidet P-40 (NP-40), 1 mM EDTA and 5% glycerol) supplemented with protease and phosphatase inhibitors (1:100 dilution) (Sigma Aldrich). Adherent cells were then scraped, transferred into pre-cooled microfuge tubes and incubated on ice for 30 min. Lysates were clarified by centrifugation at 13,300×g at 4°C for 10 min. The protein extract concentration was measured using a bicinchoninic acid assay (BCA) (Thermo Fisher Scientific) according to the manufacturer’s instructions. Proteins were resolved by electrophoresis on 20% SDS-PAGE gels. Following electrophoresis, proteins were transferred onto nitrocellulose membranes (Bio-Rad). Membranes were blocked in 1× Tris-buffered saline (TBS; Severn Biotech) containing 0.001% Tween 20 (Acros Organics) and 5% skimmed milk powder (Sainsbury’s). After washing once with TBST, membranes were incubated with primary antibody (Table 3) with gentle agitation overnight at 4°C. The following day, membranes were washed 3 times for 5 min with TBST. Membranes were subsequently incubated with rabbit or mouse secondary antibody (Thermo Fisher Scientific) for 1 h at room temperature and then washed 6 times for 5 min with TBST. Finally, the proteins were detected using Immobilon western Chemiluminescent HRP Substrate (Merck Millipore) and developed with an Odyssey Fc Imaging System (LI-COR). Human α-actin was used as a loading control.

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Table 3. Detection antibodies for immune signaling proteins.

https://doi.org/10.1371/journal.ppat.1012031.t003

Statistical analyses

All experiments were performed with at least three biological replicates. Data were analyzed using Prism 9.4 (GraphPad Software). All data used to generate the graphs is provided as source data (S3 Table).