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Molecular Biology of the Cell
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Articles
Sur7 mediates a novel pathway for PI4,5P2 regulation in C. albicans that promotes stress resistance and cell wall morphogenesis
Lanze Carla E. a
Konopka James B. a *
a Department of Microbiology and Immunology, Stony Brook University, Stony Brook, NY 11794-5222
Gladfelter Amy Susanne Monitoring Editor
Duke University
Public Health Service

National Institutes of Health (R01AI047837

National Institutes of Health training grant (NIH T32AI007539)

National Institute of Allergy and Infectious Diseases

*Address correspondence to: James B. Konopka (james.konopka@stonybrook.edu).
01 7 2024
01 7 2024
35 7 ar9923 8 2023
01 5 2024
15 5 2024
© 2024 Lanze and Konopka. “ASCB®,” “The American Society for Cell Biology®,” and “Molecular Biology of the Cell®” are registered trademarks of The American Society for Cell Biology.
2024
https://creativecommons.org/licenses/by-nc-sa/4.0/ This article is distributed by The American Society for Cell Biology under license from the author(s). It is available to the public under an Attribution 4.0 International Creative Commons CC-BY 4.0 License.

The human fungal pathogen Candida albicans can cause lethal systemic infections due to its ability to resist stress from the host and to undergo invasive hyphal growth. Previous studies showed that plasma membrane MCC/eisosome domains were important for virulence by promoting the ability of Sur7 to mediate normal cell wall morphogenesis and stress resistance. The sur7Δ mutant displayed abnormal clusters of PI4,5P2, suggesting that misregulation of this lipid underlies the sur7Δ phenotype. To test this, we increased PI4,5P2 levels by deleting combinations of the three PI4,5P2 5′ phosphatase genes (INP51, INP52, and INP54) and found that some combinations, such as inp51Δ inp52Δ, gave phenotypes similar the sur7Δ mutant. In contrast, deleting one copy of MSS4, the gene that encodes the 5′ kinase needed to create PI4,5P2, reduced the abnormal PI4,5P2 clusters and also decreased the abnormal cell wall and stress sensitive phenotypes of the sur7Δ mutant. Additional studies support a model that the abnormal PI4,5P2 patches recruit septin proteins, which in turn promote aberrant cell wall growth. These results identify Sur7 as a novel regulator of PI4,5P2 and highlight the critical role of PI4,5P2 in the regulation of C. albicans virulence properties.

Plasma membrane MCC/eisosome domains enable Sur7 to mediate normal cell wall morphogenesis and stress resistance in C. albicans. It was previously unknown how Sur7 could regulate such broad cellular effects.

Phenotypes similar to loss of SUR7 were caused by mutations that increased PI4,5P2 levels and sur7Δ mutant phenotypes were reduced by a mutation that decreased PI4,5P2 levels. Septin proteins were identified as responsible for aberrant cell wall growth.

These results identify Sur7 as a novel regulator of PI4,5P2 and highlight the critical role of PI4,5P2 in the regulation of C. albicans virulence properties.
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pmcINTRODUCTION

Candida albicans is distinct from most other fungi in that it resides in close association with humans. It typically forms a commensal association with the human skin and mucosa (Richardson et al., 2019). However, when conditions change, such as a decrease in host immunity, C. albicans can cause severe morbidity from mucosal diseases including vulvovaginal candidiasis (Rosati et al., 2020) and oral candidiasis (Vila et al., 2020). C. albicans can also cause lethal systemic candidiasis, which results from C. albicans invading the bloodstream and disseminating to different sites in the host (Lopes and Lionakis, 2022). The ability to undergo morphological transitions allows C. albicans to cause such destructive disease, as the yeast form is advantageous for bloodstream dissemination and gut colonization, whereas switching to hyphal growth allows for the penetration of epithelial cells, biofilm formation, organ colonization, and enhances phagosomal escape (Wächtler et al., 2012; Noble et al., 2017; Austermeier et al., 2020; Min et al., 2020). Other factors that make C. albicans such a successful pathogen include its ability to sense and respond to the host stress factors of elevated temperature, pH changes, nutrient availability, oxidative stress, and osmotic stress; and in some cases C. albicans can evolve resistance to antifungal drugs (Shapiro and Cowen, 2012; Dantas Ada et al., 2015; Witchley et al., 2019; Du et al., 2020; Lee et al., 2021; Lopes and Lionakis, 2022; Yaakoub et al., 2022).

The plasma membrane (PM) of C. albicans plays key roles in both morphogenesis and stress resistance. Proteins found in the PM are responsible for sensing a dynamic environment and the PM is directly involved in the secretion of virulence factors and antioxidants (Douglas and Konopka, 2016). The importance of the PM is highlighted by the fact that the three major classes of antifungal drugs directly or indirectly target PM function (Odds et al., 2003). Additionally, the PM contains proteins needed for proper morphogenesis and morphological transitions (Ibe and Munro, 2021). Recent research has illuminated the role of the Membrane Compartment of Can1 (MCC), also referred to as eisosomes, in fungal morphogenesis and stress resistance (Lanze et al., 2020). Eisosomes are stable invaginations of the PM found across the kingdom Fungi (Lee et al., 2015). In Saccharomyces cerevisiae and C. albicans they are on average 50-nm deep, 50-nm wide, and 200–300 nm long (Strádalová et al., 2009; Wang et al., 2016; Lanze et al., 2020). These membrane invaginations are stabilized at the base by Bin/amphiphysin/Rvs (BAR) domain containing proteins, Pil1 and Lsp1, that promote membrane curvature (Olivera-Couto et al., 2011; Ziółkowska et al., 2011; Zhao et al., 2013; Kabeche et al., 2015). C. albicans eisosome mutants lacking both Pil1 and Lsp1, or Sur7 alone, were shown to have strikingly abnormal cell wall invaginations and tubes of cell wall material protruding into the cell. Further research implicated Sur7 as the key eisosome protein responsible for proper morphogenesis in C. albicans (Wang et al., 2016). C. albicans cells lacking Sur7 also show increased sensitivity to cell wall and PM stress (Wang et al., 2016) and defects in invasive growth and virulence (Douglas et al., 2012).

Although its function is important, the cellular role of Sur7 is not well understood. Sur7 is a tetraspan protein that displays some amino acid similarity to the claudin family of proteins in animals (Alvarez et al., 2009) that are important for forming tight junctions, epithelial cell tubes, and lumen formation in organs (Baumholtz et al., 2017). Recent studies indicate that Sur7 regulates an intracellular function, based on the discovery that the cytoplasmic C terminus of Sur7 was necessary for promoting stress resistance and proper morphogenesis (Lanze et al., 2021). Additional studies suggested that misregulation of PI4,5P2 might contribute to the abnormal phenotypes of sur7Δ cells. In particular, the striking phenotypes of sur7Δ mutants correlated with mislocalization of a PI4,5P2 specific probe (PLCδ1 PH domain fused to RFP) to atypical patches in the PM (Wang et al., 2016). PI4,5P2 is found in small quantities on the inner leaflet of the PM of eukaryotic cells where it recruits proteins to the PM that are involved in key cellular processes (Mandal, 2020). Interestingly, septin proteins, which are known to bind PI4,5P2, mislocalized to sites of abnormal PI4,5P2 patches and cell wall growth in sur7Δ cells (Wang et al., 2016). PI4,5P2 is enriched at the bud neck in C. albicans (Vernay et al., 2012), indicating that PI4,5P2 enrichment normally recruits septins to specific sites in the cell. The ability of the septins to act as a scaffold at the bud neck to coordinate the cell wall machinery, including chitin synthase, to form the septum between dividing cells (Roh et al., 2002; Bertin et al., 2010), suggests that the septins may promote the aberrant cell wall growth at these ectopic sites in sur7Δ cells by recruiting the cell wall synthesis machinery.

Genetic studies with S. cerevisiae and Schizosaccharomyces pombe have linked eisosomes to the regulation of PI4,5P2. These results suggested a specific interaction between eisosome proteins and Inp51, a 5′ phosphatase that converts PI4,5P2 into PI4P (Fröhlich et al., 2014; Kabeche et al., 2014). The phosphorylated forms of phosphatidylinositol (PI, PI4P, and PI4,5P2) are known to recruit different proteins and activate different signaling pathways so that the phosphorylation and dephosphorylation of phosphatidylinositol species is a dynamic process active at the PM (Strahl and Thorner, 2007). PI4,5P2 5′ phosphatases have not been thoroughly characterized in C. albicans, except for Inp51 which has been studied in relation to its response to caspofungin, cell wall stress, and virulence (Badrane et al., 2008, 2012, 2016; Wang et al., 2016). Based on sequence similarity to S. cerevisiae, we identified three putative phosphatidylinositol 5′ phosphatase genes in C. albicans (INP51, INP52, and INP54) and compared the phenotypes of cells lacking different combinations of these phosphatase genes to sur7Δ cells. Altogether, these results provide the first characterization of all the PI4,5P2 5′ phosphatase genes in C. albicans and identify Sur7 as a novel regulator of PI4,5P2.

RESULTS

Sur7 and PI 4,5P 2 5′ phosphatases similarly regulate cell wall stress resistance

To test the role of PI4,5P2 in the sur7Δ phenotype, we began by deleting the genes coding for the PI4,5P2 5′ phosphatases in C. albicans. There are three predicted 5′ PI4,5P2 phosphatases in C. albicans, Inp51, Inp52, and Inp54, which share homology to the S. cerevisiae PI4,5P2 phosphatases Inp51, Inp52, Inp53, and Inp54 and the H. sapiens 4′ and 5′ PI4,5P2 phosphatases Synj1, Synj2, and Inpp5b. In S. cerevisiae, Inp52 has a paralog, Inp53, that arose from the whole genome duplication which is absent from C. albicans (Strahl and Thorner, 2007). The conservation of the protein sequence between the C. albicans proteins and the S. cerevisiae and H. sapiens homologues are detailed in Figure 1A. All three of the C. albicans Inp proteins have domains with homology to 5′ PI4,5P2 phosphatases, with Inp51 and Inp52 being more conserved (Hughes et al., 2000; Strahl and Thorner, 2007). Inp51 and Inp52 also contain 4′ phosphatase domains. However, the S. cerevisiae Inp51 is thought to be inactive due to the lack of the CX5R(T/S) motif responsible for 4′ phosphatase activity (Stolz et al., 1998; Guo et al., 1999). This motif is also absent from Inp51 in C. albicans, so this 4′ phosphatase domain is likely also inactive. The motifs implicated in catalysis for the 5′ phosphatase domains are conserved in C. albicans, as will be described further below.

FIGURE 1: Sur7 and PI4,5P2 5′ phosphatases similarly regulate cell wall stress resistance. (A) An overview of the structure of the three PI4,5P2 phosphatase proteins in C. albicans. Below the protein structures is shown the relative sequence identity to the corresponding domains of the S. cerevisiae and H. sapiens homologues. (B) Dilutions of strains indicated on the left were spotted onto agar plates under the conditions indicated on the top. The agar plates contained YPD + 35 μg/ml Calcofluor White, 35 μg/ml Congo Red, or 100 μg/ml SDS. The results are representative of six independent assays, each done in duplicate. The strains used in this study are described in the supplemental material (Supplemental Table S1).

To determine whether increasing PI4,5P2 levels results in morphogenesis and stress resistance phenotypes that are similar to the sur7Δ mutant, the inpΔ mutants were tested for susceptibility to a set of cell wall stress agents to which the sur7Δ mutant is more sensitive: Calcofluor White, Congo Red, sodium dodecyl sulfate (SDS), and high temperature (42°C) stress (Douglas et al., 2012; Wang et al., 2016; Lanze et al., 2021). We tested single inpΔ mutants (inp51Δ, inp52Δ, and inp54Δ), double inpΔ mutants (inp51Δ inp52Δ, inp51Δ inp54Δ, and inp52Δ inp54Δ), and a triple mutant lacking all three INP genes (inp51Δ inp52Δ inp54Δ). All of these inpΔ mutants represent homozygous deletions of both alleles from the diploid C. albicans genome, but for simplicity, only one delta symbol is shown in the genotypes.

The mutants lacking just one of the three INP genes generally had weaker responses to cell wall stress agents than sur7Δ (Figure 1B.) The INP gene deletion mutants lacking two out of three INP genes displayed phenotypes more comparable to sur7Δ, especially those that lacked INP52. The results also indicated that Inp52 is the most crucial PI4,5P2 phosphatase in C. albicans, and that Inp54 had the least contribution to resisting cell wall stress. The triple mutant lacking all three PI4,5P2 phosphatases was more sensitive than sur7Δ to all cell wall stress conditions. However, some of the stress sensitive phenotypes of the triple mutant can be attributed to its growth defect, as seen on the control plates grown at 37°C. Control studies showed that the cell wall stress phenotypes were largely abolished when one copy of either INP51 or INP52 was reintroduced to complement the double and triple INP mutants.

Altogether, these results provide support that the altered PI4,5P2 seen in the sur7Δ mutant underlies its sensitivity to cell wall stress, as deletion of the INP genes correlated with increased sensitivity to cell wall stress. The data also suggest that if Sur7 plays a regulatory role for the PI4,5P2 phosphatases, it likely regulates more than one of the Inp proteins, but does not fully regulate all three of them.

Sur7 and PI 4,5P 2 5′ phosphatases similarly regulate PM stress resistance.

We next evaluated the ability of strains lacking the PI4,5P2 5′ phosphatases to resist stress from copper and duramycin, as the sur7Δ mutant has previously exhibited increased sensitivity to these agents (Douglas and Konopka, 2019). Copper is thought to act by binding the lipid phosphatidylserine with high affinity (Monson et al., 2012), and duramycin by binding to the lipid phosphatidylethanolamine (Hullin-Matsuda et al., 2016). Both phosphatidylserine and phosphatidylethanolamine are normally enriched on the inner leaflet of the PM (Pomorski and Menon, 2016), therefore increased sensitivity to duramycin and/or copper indicate that sur7Δ possesses an altered PM architecture. Thus, we wanted to determine whether increasing PI4,5P2 levels by deleting the PI4,5P2 5′ phosphatases would alter PM architecture similar to the sur7Δ mutant.

Using disk diffusion assays, we found that the inpΔ mutants had increased sensitivity to both CuSO4 and duramycin similar to sur7Δ cells. All of the mutants tested had increased sensitivity to copper when compared with WT cells, and the mutants containing inp52Δ had statistically significant differences (Figure 2A). In response to duramycin, the sur7Δ mutant and all of the inpΔ mutants except for inp54Δ displayed increased sensitivity. However, the only statistically significant increases were found in the double deletion mutants containing inp52Δ and the triple deletion mutant (Figure 2B). Control studies showed that the mutant phenotypes were rescued by genetic complementation. These results indicate that the sur7Δ and inpΔ mutants possess similarly altered PM architecture, and further implicate Inp52 as the key PI4,5P2 phosphatase in C. albicans.

FIGURE 2: Sur7 and PI4,5P2 5′ phosphatases similarly regulate PM stress resistance. The susceptibility of sur7Δ and inpΔ mutants to (A) copper (CuSO4) and (B) duramycin was determined by disk diffusion halo assays in which 1 × 107 cells of the indicated strain were spread onto the surface of synthetic medium agar plates. Filter discs containing different amounts of the indicated compounds were placed onto each plate. The zone of growth inhibition was recorded in millimeters and results are presented as the change in the zone of growth inhibition relative to the wild type control strain. The results represent that average of three independent experiments and were analyzed by one way ANOVA (* < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001).

Sur7 and PI 4,5P 2 5′ phosphatases similarly regulate cell wall growth.

The sur7Δ mutant has been previously shown to have a strikingly abnormal cell wall (Alvarez et al., 2008; Wang et al., 2016). To determine whether excess PI4,5P2 underlies these cell wall abnormalities, we visualized cell wall chitin by Calcofluor White staining and found that some of the inpΔ mutants had abnormal deposits of chitin, similar to the sur7Δ mutant (Figure 3A). The inp51Δ and inp52Δ mutants had moderate abnormalities, with small patches of abnormal chitin staining in some of the cell population. The double inpΔ mutants lacking INP52 and the triple deletion mutant displayed larger areas of abnormal cell wall, similar to sur7Δ (Figure 3A). The extent of the chitin abnormalities was quantified in Figure 3B. All of the mutants except inp54Δ had statistically significant differences in chitin abnormalities when compared with the WT control strain, and the triple inpΔ mutant had the most similar percentage of cells displaying Calcofluor White staining abnormalities to sur7Δ.

FIGURE 3: Sur7 and PI4,5P2 phosphatases similarly regulate cell wall growth. (A) C. albicans sur7Δ and inpΔ mutant strains were stained with Calcofluor White to reveal abnormal invaginations of cell wall growth, some of which are specified by white arrows. The scale bar indicates 3 µm. (B) A bar graph representing the percentage of chitin abnormalities for strains shown in panel A. The results represent the average of three independent assays and were analyzed by one way ANOVA for their difference to the WT control strain (*** < 0.001, **** < 0.0001). (C) Cell sections of sur7Δ and inpΔ double and triple mutant strains were captured by transmission electron microscopy. The sur7Δ, inp51Δ inp52Δ, and inp51Δ inp52Δ inp54Δ mutants are characterized by thicker cell walls and invaginations of cell wall. The black arrows indicate regions where there are spiky invaginations, while white arrows point to the cell wall tubes. The scale bar indicates 500 nm. (D) Average cell wall thickness of the indicated strains detected by transmission electron microscopy. Values were analyzed by one way ANOVA for their difference to the WT control strain (**** < 0.0001).

To further assess the abnormal cell wall phenotype, we employed transmission electron microscopy. As expected from previous studies (Alvarez et al., 2008), the sur7Δ mutant was characterized by a thickened cell wall, invaginations of cell wall material, and inwardly protruding tubes of cell wall material (Figure 3C). Interestingly, the inp51Δ inp52Δ mutant displayed cell wall abnormalities that were very similar to sur7Δ cells. The inp51Δ inp54Δ mutant appeared to have no major cell wall defects, while inp52Δ inp54Δ had moderate defects. Remarkably, the inp51Δ inp52Δ inp54Δ triple mutant had a severe cell wall defect characterized by an extremely thick cell wall and multiple cell wall invaginations. The cell wall thickness, as observed by transmission electron microscopy, is quantified in Figure 3D. These data further support our hypothesis that the sur7Δ phenotype is caused by abnormal PI4,5P2 regulation and that if Sur7 regulates the Inp proteins it must regulate more than one, but not all of them.

Sur7 and Inps regulate hyphal formation and invasive growth.

Mutants lacking Sur7 have been shown previously to be defective at hyphal growth in liquid media and invasive growth into agar (Wang et al., 2016), consistent with a defect in invasive hyphal growth in vivo (Douglas et al., 2012), which is an important virulence factor for C. albicans (Min et al., 2020). We therefore examined the inpΔ mutants to assess whether abnormal PI4,5P2 regulation underlies these sur7Δ growth defects. The sur7Δ mutant typically fails to make hyphae when induced or makes short wide hyphae (pseudohyphae), and only occasionally makes relatively normal hyphae. The single and double inpΔ mutants lacking Inp52 displayed a similar hyphal defect to sur7Δ. Mutants lacking Inp51 sometimes displayed slightly thickened hyphal filaments. The mutant lacking all three INP genes had the most severe defect in forming hyphae, with most of the cells showing no signs of initiating hyphal formation. Mutants complemented with one copy of INP52 largely restored hyphal growth (Figure 4, A and B).

FIGURE 4: Sur7 and Inps regulate hyphal and invasive growth. (A) The sur7Δ and inpΔ mutant strains were tested for their ability to initiate hyphal growth. Cells were grown overnight at 30°C in synthetic media and inoculated into fresh synthetic media containing 50 mM GlcNAc. Cultures were subsequently incubated at 37°C for 2.5 h to induce hyphal formation. The scale bar indicates 3 µm. (B) A bar graph representing the quantification of the cell morphologies displayed above in panel A. The number of cells in hyphal, pseudohyphal, and nonfilamentous form were counted. The results represent the average of three independent assays. (C) The indicated strains were tested for their ability to grow invasively into plates containing 1.5 agar and 4% bovine serum. The cells were incubated at 37°C for 7 d and then photographed. The results represent six independent assays, each done in duplicate.

A similar pattern was observed for invasive growth, with sur7Δ and inp52Δ having comparable defects. The double deletion mutants lacking Inp52 and the mutant lacking all three INP genes had slightly more severe defects than sur7Δ in invasive growth. The inp51Δ and inp54Δ mutants were only slightly defective at invasive growth, showing shorter hyphal radiation than in the WT strain. These results were in general agreement with a previous analysis of inp51Δ (Badrane et al., 2008). All of the defects were reversed by complementing one copy of the appropriate gene (Figure 4C) These data support the hypothesis that increased PI4,5P2 underlies the invasive and hyphal growth defects of sur7Δ, and that Inp52 is this most critical of the PI4,5P2 5′ phosphatases.

The sur7Δ and inpΔ mutants have mislocalized PI 4,5P 2 with abnormal PM structures and ectopic septins

To visualize PI4,5P2 in the sur7Δ and inpΔ mutants, a previously described probe containing the human PLCδ1 PH domain fused to RFP was utilized (Badrane et al., 2012). This probe appeared to stain the PM in inp51Δ, inp54Δ, and inp51Δ inp54Δ strains similarly to the WT control cells. In contrast, the sur7Δ mutant and mutants lacking Inp52 contained abnormal patches of the PH domain probe. Interestingly, it was difficult to visualize the RFP signal from the PM in the inp51Δ inp52Δ and inp52Δ inp54Δ mutants, possibly because the strong signal from the PI4,5P2 patches was sequestering more of the probe. To confirm that the PI4,5P2 patches corresponded to excess PM, we GFP tagged Pma1, a major PM protein (Athanasopoulos et al., 2019). In the sur7Δ, inp52Δ, inp51Δ inp52Δ, and inp52Δ inp54Δ mutants, the abnormal PI4,5P2 patches colocalized with Pma1-GFP. The inp51Δ inp52Δ mutant was also characterized by sometimes displaying an irregularly shaped PM that surrounded the abnormal PI4,5P2 patches (Figure 5A). The percentage of cells displaying abnormal Pma1-GFP is quantified in Figure 5C. These results indicate that deletion of the PI4,5P2 phosphatases causes aberrant PM invaginations that colocalize with patches of the PI4,5P2 probe similar to what was seen in the sur7Δ mutant.

FIGURE 5: The sur7Δ and inpΔ mutants have mislocalized PI4,5P2 with abnormal PM structures and ectopic septins. Fluorescence microscope images of the indicated sur7Δ and inpΔ mutant strains that contain both a PLCδ1 PH domain fused to RFP (for visualizing PI4,5P2, top row) and a GFP tag of either (A) Pma1 or (B) Cdc10 (2nd row). The 3rd row contains the merged image, and the bottom rows are the same cells photographed using DIC optics. The scale bar indicates 3 µm. (C) Bar graph summarizing the percentage of cells displaying abnormal Pma1 structures for strains in panel A. (D) Bar graph summarizing the percentage of cells displaying abnormal Cdc10 localization for strains in panel B. The results represent the average of three independent assays and were analyzed by one way ANOVA for their difference to the WT control strain (* < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001).

The sur7Δ mutant was previously shown to display abnormal localization of the septin protein Cdc10, which colocalized with abnormal PI4,5P2 patches, consistent with previous reports that the septin proteins bind PI4,5P2 (Bertin et al., 2010). To assess whether an excess of PI4,5P2 causes the mislocalization of septin proteins, we examined Cdc10-GFP in the inpΔ mutants. In the WT and inp54Δ mutant strains, Cdc10 localized properly to bud necks. Conversely, the other mutants contained Cdc10-GFP that localized to sites of abnormal PI4,5P2 patches with sur7Δ, inp51Δ inp52Δ, and inp52Δ inp54Δ containing similarly high percentages of cells with abnormal Cdc10-GFP localization (Figure 5, B and D). Septin proteins coordinate cell wall building machinery and are needed for proper chitin deposition (DeMarini et al., 1997; Oh and Bi, 2011). Thus, recruitment of septins to abnormal PI4,5P2 patches may drive the abnormal cell wall morphology of sur7Δ. Altogether, the results show that the deletion of INP genes, especially INP52, correlates with abnormal invaginations of the PM, altered PI4,5P2, and mislocalized septins.

The 5′-phosphatase domain of Inp52 regulates PI 4,5P 2 and cell wall morphology

The evidence indicates that Inp52 is the most crucial PI4,5P2 phosphatase in C. albicans, thus we aimed to determine which part of this protein is responsible for its activity. Inp52 is distinct from Inp51 and Inp54 in that it is predicted to have both a functional 4′ and a 5′ phosphatase domain (Strahl and Thorner, 2007). The catalytic motifs of the Inp proteins are conserved up to mammalian PI4,5P2 phosphatases (Guo et al., 1999; O’Malley et al., 2001) so we chose the three most highly conserved motifs for our mutagenic analysis (Whisstock et al., 2002). For the underlined catalytic residues shown in Figure 6A, we mutated those residues to alanine and named the strains containing these mutants INP52-4p (key residue mutation in the 4′-phosphatase domain [D435A]), INP52-5p,1 (key residue mutation in Motif 1 of the 5′-phosphatase domain [D767A]), and INP52-5p,2 (key residue mutation in Motif 2 of the 5′-phosphatase domain [D843A.])

FIGURE 6: The 5′-phosphatase domain of Inp52 regulates PI4,5P2 and cell wall morphology. (A) A schematic showing the three regions of Inp52 that were chosen for mutational analysis. The three most conserved catalytic motifs were chosen, one is in the 4′ phosphatase domain and two are in the 5′ phosphatase domain. The catalytic residues chosen for mutation to alanine are in bold and underlined. Amino acid sequences from C. albicans, S. cerevisiae, and H. sapiens are shown to highlight the conserved amino acids. (B) Distribution of the PLCδ1 PH-RFP probe (for visualizing PI4,5P2) in inp52Δ, the complemented strain, and all three of the catalytic site mutants (bottom panel) with DIC optics (top panel). (C) Bar graph summarizing the percentage of PI4,5P2 abnormalities for strains in panel B. The results represent the average of three independent assays and were analyzed by one way ANOVA for their difference to the INP52 complemented strain (* < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001). (D) Spot assays of catalytic mutants in the inp51Δ inp52Δ strain background (top panel) and the inp52Δ inp54Δ strain background (bottom panel.) Dilutions of indicated strains were spotted onto agar plates under the conditions indicated on the top. The agar plates contained YPD + 35 μg/ml Calcofluor White, 35 μg/ml Congo Red, or 100 μg/ml SDS. The results are representative of three independent assays, each done in duplicate. (E) The inp52Δ catalytic mutant strains were tested for their ability to grow invasively (top panel) and form hyphae (bottom panel). To test for invasive growth, strains were spotted onto plates containing 1.5% agar and 4% bovine serum. The plates were incubated at 37°C for 7 d and then photographed. To test for hyphal induction, cells were grown overnight at 30°C in synthetic media and inoculated into fresh synthetic media containing 50 mM GlcNAc. Cultures were subsequently incubated at 37°C for 2.5 h to induce hyphal formation. (F) A bar graph representing the quantification of the cell morphologies displayed above in panel E (bottom). The number of cells in hyphal, pseudohyphal, and nonfilamentous form were counted. The results represent the average of three independent assays. The scale bars indicate 3 µm.

The wild type INP52 and the three mutant versions were then introduced into an inp52Δ mutant strain to test their function. As expected, control studies showed that inp52Δ mutant displayed abnormal patches of the PLCδ1 PH domain-RFP probe whereas reintroduction of wild-type INP52 restored a normal cortical appearance of the probe. Similar results were observed for the inp52Δ mutant transformed with mutations in the 4′-phosphatase domain and Motif 2 in the 5′-phosphatase domain, both of which resembled inp52Δ mutants complemented with one copy of INP52. In contrast, the inp52Δ mutant transformed with one copy of INP52 containing the mutation in Motif 1 of the 5-phosphatase domain displayed abnormal PI4,5P2 patches as seen in the mutant completely lacking INP52 (Figure 6, B and C) This implicates Motif 1 of the 5′ phosphatase domain as being indispensable for Inp52 activity.

To strengthen these results, and further link PI4,5P2 phosphatase activity to facilitating proper cell wall morphogenesis, we performed spot assays of inp52Δ catalytic mutants to cell wall stress agents. As seen in Figure 1, the inp51Δ inp52Δ and inp52Δ inp54Δ mutants are very sensitive to cell wall stress agents, thus these were chosen as the background strains. In both backgrounds, complementing with the INP52-5p,1 mutation failed to restore stress resistance, especially to Calcofluor White, SDS, and high temperature, further implicating Motif 1 as critical for Inp52 activity. In the inp52Δ inp54Δ background, complementing with INP52-4p and INP52-5p,2 mutant genes fully restored all the stress resistance phenotypes, except slightly less for Calcofluor White. Complementing inp51Δ inp52Δ with the INP52-4p and INP52-5p,2 mutant genes restored its resistance to high temperature stress and Congo Red, but less so for Calcofluor White and SDS (Figure 6D).

A similar pattern followed when this mutant set was tested for ability to form hyphae and grow invasively. Introducing the INP52-4p mutant gene caused no defects for invasive and hyphal growth and the INP52-5p,2 mutant gene caused slight defects. In contrast, strains that received the INP52-5p,1 mutant gene were completely defective at invasive and hyphal growth (Figure 6, E and F). This indicates that the 5′-phosphatase activity of Inp52 is a major regulator of PI4,5P2 and cell wall morphology.

To examine whether Inp52 is the most crucial of the 5′ PI4,5P2 phosphatases because of its functional properties or because it is produced at higher levels than Inp51 and Inp54, we probed the levels of the Inp proteins using Western blotting. Inp54 showed the highest signal on the blots (Supplemental Figure S1A), and Inp51 showed only slightly lower level than Inp52 (< twofold), indicating that Inp52 has special properties. Analysis of the catalytic mutant proteins was examined to determine whether their phenotypes were due to loss of function or altered protein production. This proved to be more challenging as the Inp proteins are low abundance and the catalytic mutant genes were only present in one copy. Although all three catalytic mutant proteins were present at lower levels, the overall the levels were not that different indicating that the inp52Δ mutant phenotypes were not due to changes in protein levels (Supplemental Figure S1B).

Increased PI 4,5P 2 levels and abnormal coordination of downstream effectors are responsible for the sur7Δ phenotype

The similar phenotypes between the inpΔ and sur7Δ mutants strongly implicated altered PI4,5P2 in the sur7Δ phenotype. To test this, we decreased the levels of PI4,5P2 in sur7Δ by deleting one copy of MSS4, the sole 5′ PI4,5P2 kinase in C. albicans that converts PI4P to PI4,5P2 (Vernay et al., 2012). Consequently, in the sur7Δ/Δ mss4Δ/MSS4 mutant, we observed a sharp reduction in abnormal PI4,5P2 patches and the severity of the abnormal PI4,5P2 phenotype (Figure 7, A and B). We similarly saw a decrease in the number of cells with chitin abnormalities in the sur7Δ/Δ mss4Δ/MSS4 mutant (Figure 7, C and D).

FIGURE 7: Increased PI4,5P2 levels and abnormal coordination of downstream effectors are responsible for the sur7Δ phenotype. (A) Distribution of the PLCδ1 PH-RFP probe (for visualizing PI4,5P2) in WT, sur7Δ, and sur7Δ mss4Δ (top panel) and with DIC optics (bottom panel). (B) Bar graph summarizing the percentage of PI4,5P2 abnormalities for strains in panel A. (C) WT, sur7Δ, and sur7Δ mss4Δ were stained with Calcofluor White to reveal abnormal invaginations of cell wall growth (top panel), the DIC optics images of the same cells are below. (D) The percentage of chitin abnormalities for strains in panel C. (E) Cell size measurements and (F) chitin abnormalities for the indicated strains. The scale bars indicate 3 µm. The results represent the average of three independent assays and were analyzed by one way ANOVA (* < 0.05, ** < 0.01, *** < 0.001, **** < 0.0001).

We next determined the role of septins and chitin synthase in the sur7Δ phenotype. Mutants lacking the septin genes CDC10 or CDC12 have previously been shown to have destabilized septin rings (Warenda and Konopka, 2002; Li et al., 2012). Thus, we predicted that disruption of these genes in the sur7Δ mutant would decrease ectopic septin localization and reduce the amount of abnormal cell wall. We made combinatorial mutants in the sur7Δ background that lack both copies of CDC10, one copy of CDC12, or one copy of CHS3 (a chitin synthase). Our results showed that these combinatorial mutants all resulted in a decrease in cell size (Figure 7E; Supplemental Figure S2A) and chitin abnormalities (Figure 7F; Supplemental Figure S2B) in the sur7Δ mutant, providing further support that septins and chitin synthase contribute to sur7Δ abnormal cell wall phenotype.

Inp proteins localize to the cytoplasm and punctate patches in the PM

To further define the cellular functions of the Inps, we employed GFP-tagging to determine their subcellular localization. We found that Inp51, Inp52, and Inp54 all displayed a few bright puncta near the cell periphery, as well as GFP signal detected in the cytoplasm (Figure 8). To determine whether these puncta colocalized with Sur7, we tagged Sur7 with mScarlet. Sur7-mScarlet displayed a characteristic localization to MCC/eisosome domains, as it was found in puncta around the cell periphery. Some, but not all of the Inp52-GFP and Inp54-GFP patches colocalized with Sur7 (Figure 8). These results suggest that Sur7 and the PI4,5P2 5′phosphatases may directly interact, as they can be found in the same subcellular location.

FIGURE 8: Inp proteins localize to the cytoplasm and punctate patches in the PM. Fluorescence microscopy images of C. albicans strains. The top row shows images of cells producing the indicated Inp51-GFP, Inp52-GFP, or Inp54-GFP. The second row shows detection of the eisosome marker Sur7-mScarlet in the same cells. The third row shows the merge. The bottom row shows DIC optics images of indicated strains. The insets on the lower right of the merge panels show tracings through the patches of Inp52-GFP and Inp54-GFP that are indicated with an arrow. The scale bar indicates 3 µm.

DISCUSSION

Sur7 plays critical roles in guiding C. albicans cell wall morphogenesis, which is important for invasive hyphal growth and avoiding immune detection (Childers et al., 2020). It also mediates resistance to a wide range of stressors, including agents that affect the cell wall and PM, such as resistance to copper that is important for survival in macrophages (Douglas et al., 2012; Festa and Thiele, 2012). Because sur7Δ cells display such diverse phenotypes, the question emerged as to how Sur7 can regulate so many different cellular properties. Our results show that Sur7 has such a broad impact because it regulates PI4,5P2, a PM lipid critical for proper cellular signaling and morphogenesis. Deletion of the INP genes to increase PI4,5P2 caused defects similar to the sur7Δ mutant (Figures 1–5). These defects were suppressed in the sur7Δ mutant by deleting one copy of MSS4, which phosphorylates PI4,P to create PI4,5P2 (Figure 7). These results identify Sur7 as a novel regulator of PI4,5P2. This is broadly significant because PI4,5P2 is a key cellular regulator in wide range of eukaryotic cells (Mandal, 2020).

Roles of Sur7 and PI 4,5P 2 in morphogenesis

The strikingly abnormal morphology of sur7Δ cells, characterized by invaginations of cell wall, often in a tubular form, was recapitulated in the inpΔ mutants. This was especially true for the inp51Δ inp52Δ mutant that was essentially indistinguishable from the sur7Δ mutant in TEM images (Figure 3). The abnormal cell wall invaginations seen in the sur7Δ and inpΔ mutants were associated with patches of PI4,5P2 detected with a PLCδ1 PH domain probe. It was interesting that the increased PI4,5P2 did not simply lead to a stronger signal evenly around the PM. Instead, the probe revealed a patchy distribution of PI4,5P2 (Figure 5), consistent with a previous report that excess PI4,5P2 clusters into patches in S. cerevisiae (Riggi et al., 2018). The PI4,5P2 patches were associated with the presence of ectopic septins, which are known to bind PI4,5P2 (Bertin et al., 2010). For example, the septin Cdc10 was mislocalized to PI4,5P2 patches in inpΔ and sur7Δ cells (Figure 5), rather than being exclusively found at the bud neck, which is enriched in PI4,5P2 (Vernay et al., 2012). Septin proteins act as a scaffold to recruit cell wall synthesis machinery needed for proper chitin deposition (DeMarini et al., 1997; Oh and Bi, 2011). Destabilizing septins in sur7Δ strain by mutating CDC10 or CDC12 reduced the abnormal cell wall phenotype, supporting a direct role for the septins in this phenotype (Figure 7). A similar reduction was seen after deleting one copy of the CHS3 chitin synthase from sur7Δ cells, consistent with a model that the PI4,5P2 patches recruit septins that promote the aberrant cell wall growth.

Some of the aberrant cell wall and PM structures observed in sur7Δ cells may possibly be due to altered eisosome structures. In this regard, a recent study showed observed unusual eisosome structures in an S. cerevisiae inp51Δ inp52Δ mutant (Haase et al., 2023). However, it was not clear whether this led to altered cell wall formation. Furthermore, the eisosome furrows formed in a C. albicans sur7Δ mutant appeared to be normally shaped when analyzed at high resolution by freeze-etch EM (Wang et al., 2016).

All three C. albicans INP genes were important for invasive hyphal growth (Figure 4), consistent with a gradient of PI4,5P2 at the hyphal tip being important for promoting highly polarized filamentous growth (Vernay et al., 2012). Thus, the abnormal distribution of PI4,5P2 in sur7Δ cells likely contributes to the hyphal defect. Furthermore, because eisosomes take time to form, and are therefore not present at the hyphal tip, this suggests a possible role for Sur7 in diminishing the PI4,5P2 signal in mother cells in order to enhance the PI4,5P2 gradient at the hyphal tip. Other roles for Sur7 in cell wall morphogenesis were suggested by the localization of Sur7 to sites where the cell wall was damaged by neutrophil attack (Hopke et al., 2016).

Sur7 regulates PM asymmetry indirectly through PI 4,5P 2

The increased sensitivity of sur7Δ to copper and duramycin was linked to PI4,5P2, as the same phenotypes were present in inpΔ mutants (Figure 2). The elevated susceptibility of sur7Δ cells to copper and duramycin indicates the abnormal presence in the outer leaflet of the PM of phosphatidylserine and phosphatidylethanolamine, respectively (Douglas and Konopka, 2019). These phospholipids are usually enriched in the inner leaflet due to the action of phospholipid flippases (Sakuragi and Nagata, 2023), which suggests that altered PI4,5P2 can impact their function (Douglas and Konopka, 2019). Other phenotypic similarities with the inpΔ mutants indicate that increased PI4,5P2 underlies additional PM phenotypes of sur7Δ, such as increased sensitivity to SDS. Thus, the results indicate that some sur7Δ phenotypes are due directly to PI4,5P2, and others are due to downstream effects resulting from elevated PI4,5P2, such as changes in phosphatidylserine and phosphatidylethanolamine. Alterations in these phospholipids is known to impact virulence (Cassilly and Reynolds, 2018). Altogether, these results further reveal how Sur7 can broadly impact cellular functions outside of the eisosomes.

Regulation of PI 4,5P 2 by Sur7 and eisosomes

Previous genetic studies with S. pombe and S. cerevisiae implicated eisosomes in regulating PI4,5P2 by acting in a pathway with INP51 (Fröhlich et al., 2014; Kabeche et al., 2014). The Inp51 protein was also shown to localize to eisosomes in S. cerevisiae (Fröhlich et al., 2014). However, the S. cerevisiae and S. pombe pil1Δ mutants are defective in forming eisosomes, which makes it difficult to conclude what Pil1 is doing directly and what it is doing through its role in forming the eisosome furrows. In C. albicans, sur7Δ and pil1Δ lsp1Δ give similar phenotypes, but Sur7 is the key regulator of PI4,5P2 because a sur7Δ mutant still forms eisosomes and overproduction of Sur7 partially rescued the defects of a pil1Δ lsp1Δ mutant (Wang et al., 2016). An interesting question for future studies is whether Sur7 affects PI4,5P2 through regulation of the Inp proteins, or indirectly through other mediators. This analysis will be complicated by the experimental difficulties of developing approaches to study eisosome furrows in vitro and the very low levels of the Inp proteins. However, consistent with a direct role, the cytoplasmic C terminus of the tetraspan Sur7 protein is a key functional domain, indicating Sur7 regulates an intracellular function (Lanze et al., 2021), GFP-tagged Inp proteins partially colocalized with Sur7 in C. albicans eisosomes (Figure 8), and Inp51 was detected at eisosomes in S. cerevisiae (Fröhlich et al., 2014). On the other hand, other proteins could be involved because >25 proteins localize to eisosomes (Lanze et al., 2020), and Inp51 requires association with Irs4 to function (Badrane et al., 2012).

Another difference between species is that in S. cerevisiae and S. pombe, pil1Δ displayed genetic interactions indicating it acts in a pathway specifically with inp51Δ (S. pombe syj1Δ) and not the other the PI4,5P2 5′ phosphatases (Fröhlich et al., 2014; Kabeche et al., 2014). Additionally, S. cerevisiae Pil1 was important for recruiting Inp51 to the eisosomes, but not other Inps (Fröhlich et al., 2014). In contrast, the C. albicans sur7Δ phenotypes are more pronounced than those of the single inp51Δ, inp52Δ, or inp54Δ mutants, suggesting that whether Sur7 acts by regulating the Inps, it must be capable of acting on more than one Inp protein. It was also distinctive that the sur7Δ phenotype is most similar inp52Δ in C. albicans. Other significant differences between species include the fact that only Pil1 is essential for eisosome formation in S. cerevisiae (Walther et al., 2006), whereas Pil1 and Lsp1 play equal roles in C. albicans (Wang et al., 2016). A further difference in the regulation of PI4,5P2 is that we found that all of the INP genes could be deleted in C. albicans, whereas in S. cerevisiae an inp51Δ inp52Δ inp53Δ mutant is lethal even though it still possesses INP54 (O’Malley et al., 2001). Similarly, in S. pombe the syj1Δ (inp51Δ) inp53Δ mutant is lethal despite also having SYJ2 (INP52; Kabeche et al., 2014).

PM invaginations and PI 4,5P 2 regulation

It is surprising that although PM eisosome furrows are present across the fungal kingdom, the function of Pil1, Lsp1 and Sur7 eisosome proteins are not highly conserved across species (see above and reviewed in [Lanze et al., 2020]) and the amino acid sequences of these proteins are not highly conserved across species (Lee et al., 2015). This raises the possibility that the membrane invaginations may be the conserved structure that is key for function, freeing the eisosome proteins to evolve in the different species. In this regard it is interesting that mammalian cells, which lack eisosomes, contain PM invaginations termed caveolae that are enriched in a pool of PI4,5P2 that is key for signaling (Pike and Casey, 1996; Morris et al., 2006; Fujita et al., 2009) and that newly described PM furrows in Caenorhabditis elegans termed meisosomes are enriched in PI4,5P2 (Aggad et al., 2023). Furthermore, the yeast bud neck, a type of membrane invagination, is enriched in PI4,5P2 (Vernay et al., 2012). These similarities support the idea that membrane invaginations are advantageous for PI4,5P2 regulation and signaling. In view of the key roles PI4,5P2 plays in both fungi and humans, and that dysregulation of PI4,5P2 is implicated in many human diseases (Mandal, 2020), further defining the mechanisms that regulate PI4,5P2 will be important to open new avenues for developing novel therapeutic strategies.

MATERIALS AND METHODS

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Growth media and strain construction

Cells were grown in rich YPD medium, which contains yeast extract, peptone, and dextrose or in synthetic medium containing yeast nitrogen base, dextrose, uridine, and amino acids for auxotrophic strains (Sherman, 1991). Supplemental Table S1 details the strains used for this study and Supplemental Table S2 lists the oligonucleotides that were used. Gene deletion mutants were made by homologous recombination and electroporation (De Backer et al., 1999), using SN152 as the starting strain (Noble and Johnson, 2005). The gene deletion mutants were made using the transient CRISPR-Cas9 system, essentially as described (Min et al., 2016). The gene-deletion constructs were synthesized using ARG4, HIS1, or the SAT1 flipper as template DNA (Reuß et al., 2004; Noble and Johnson, 2005). The primers included 20 bases of homology to these selectable marker cassettes and 80 base pairs of homology directly upstream or downstream of the targeted open reading frame (ORF). For GFPγ (Zhang and Konopka, 2010) and mScarlet (Frazer et al., 2019) tagging of proteins, the same transformation methods were utilized. The forward primer contained 80 bases of homology to the gene right before the stop codon and 20 bases of homology to the appropriate plasmid containing the fluorescent protein and selectable marker. The reverse primer contained 80 bases of homology to the region around 50 bases after the targeted ORF and 20 bases of homology to the appropriate plasmid containing the fluorescent protein and selectable marker, as previously described (Zhang and Konopka, 2010).

To assess the genotype of the transformants, genomic DNA was isolated by phenol-chloroform DNA extraction and then used as a template for several different PCR analyses to confirm the identity. The PCR primers flanked the outside and inside of the targeted ORF, as well as tested for the presence of the appropriate selectable marker in the proper location. To assess the fluorescent protein tags, forward primers were in the appropriate ORF and reverse primers were in the selectable marker genes. For the transformants containing the SAT1 flipper cassette, transformants were selected on YPD plus 200 μg/ml nourseothricin (NAT; Gold Biotechnology). To excise SAT1 from mutants, colonies were streaked onto maltose agar medium (1% yeast extract, 2% peptone, 2% maltose) and then several colonies were streaked onto YPD plus NAT plates to test for resistance or sensitivity.

Strains were made prototrophic by PCR-amplifying C. albicans ARG4, LEU2, or HIS1 with primers containing 20 bases of homology located roughly 500 base pairs upstream and downstream of the ORFs. These PCR fragments were purified with the QIAquick PCR Purification Kit (Qiagen, Germantown, MD) and transformed into their native loci. Gene complementation was achieved by integration into the Neut5L locus, as previously described (Gerami-Nejad et al., 2013). Plasmids were made by designing primers with 30 base pairs of homology to the complementing gene and 70 pairs of homology to the pDIS3 plasmid (Gerami-Nejad et al., 2013) and gap repair cloning the PCR fragments into the S. cerevisiae strain L40. Plasmids were then transformed into the Escherichia coli strain DH5α by electroporation. Resulting plasmids were digested with SfiI (New England Biolabs) and the fragments were used for transformation. This same methodology was employed to make the catalytic mutants of INP52. PCR of INP52 was undertaken using primers containing the desired base pair changes. For creating catalytic mutants fused to GFP, strains of Inp-GFP fusions were used as the DNA in the PCR reaction and a reverse primer was used to amplify the end of GFP. The presence of the appropriate mutations in the resulting mutated inp52 was confirmed by DNA sequencing. New Neut5L vectors were created as previously described (Gerami-Nejad et al., 2013) by PCR amplifying selectable markers LEU2 and HIS1 from plasmids containing homology to the Neut5L vector pDIS2 (Gerami-Nejad et al., 2013) and using gap repair in S. cerevisiae. These new vectors were needed for strains containing the PH domain probe, as those and pDIS2 both contain nonrecyclable NAT markers.

Western blot analysis

Western blots were performed to compare levels of the Inp proteins and confirm production of the catalytic mutant proteins. Strains were grown in YPD at 30°C with shaking until a density of ∼1 × 107 cells/ml was reached. Cells were harvested by centrifugation and pellets were frozen at –80°C. Cell pellets were lysed in 1x Laemmli buffer (2% SDS, 10% glycerol, 125 mM Tris-HCl, pH 6.8) by agitation with zirconia beads for one minute followed by a 1-min ice bath (four rounds total.) The insoluble fraction was removed by centrifugation, 2-mercaptoethanol was added to 5% final concentration and bromophenol blue to 0.002% final concentration, and then samples were heated at 100°C for 10 m. The proteins were separated by electrophoresis on an 8% acrylamide gel, then transferred to a 0.2 µm nitrocellulose membrane (Amersham Protran, catalogue# 10600032) using a semidry transfer apparatus. Blots were probed with a mix of two anti-GFP monoclonal antibodies (catalogue# 632381 JL-8 and catalogue# 632569 EGFP, Takara Bio, San Jose, CA) at a 1:1,000 dilution in TBS-T buffer (0.1% Tween-20, 2% [wt/vol] bovine serum albumin [BSA], and 0.2% [wt/vol] sodium azide) for 12 h. The blots were washed in TBS-T buffer and then incubated with antirabbit IgG secondary antibody (IRDye 800–conjugated, LI-COR Biosciences, Lincoln, NE) and diluted 1:10,000 in Tris-buffered saline containing 0.3% Tween-20. For Coomassie-stained gels, following SDS–PAGE, gels were stained in a Coomassie brilliant blue solution (0.1% Coomassie R-250, 40% ethanol, 10% acetic acid) for 4 h and were destained overnight with a solution of 40% methanol and 10% acetic acid. Blots were scanned with an Odyssey CLx Infrared Imaging System (LI-COR Biosciences) and the images were analyzed using ImageStudio software (LI-COR Biosciences).

Protein sequence alignments

The Inp protein sequences were obtained from the S. cerevisiae Genome Database (www.yeastgenome.org) and NCBI BLAST (www.ncbi.nlm.nih.gov/BLAST/) searches were carried out to identify homologues in the C. albicans Genome Database (www.candidagenome.org).

Microscopic analysis

Cells were grown overnight in rich or synthetic media to log phase and then analyzed using a Zeiss Axio Observer 7 microscope. Pictures were obtained with a Zeiss AxioCam 702 digital camera. Zeiss ZEN software was used to quantify cell size, create a colocalization profile, and for deconvolving images. Deconvolution of images was performed with three Z-axis images of the midsection of the cell. DIC optics were used to capture brightfield images of cells. To induce hyphal formation cells were grown overnight at 30°C in synthetic media and inoculated into fresh synthetic media containing 50 mM GlcNAc. Cultures were subsequently incubated at 37°C on a tube roller for 2.5 h and analyzed by microscopy. To visualize chitin, cells were stained as previously described using 20 μg/ml Calcofluor White (Fluorescent Brightener 28; Sigma-Aldrich, St. Louis, MO; Pringle, 1991). This technique was modified by washing the cells with methanol and acetone as previously described (Wang et al., 2016) to remove the PM and allow the dye to access the invaginations of cell wall material.

Cells were prepared for transmission electron microscopy by growing overnight at 30°C to log phase in YPD. The cells were then fixed in a solution of 0.1M sodium cacodylate buffer (pH 7.4) with 3% EM-grade glutaraldehyde at room temperature for 1 h. After fixation, cells were rinsed three times in 0.1M sodium cacodylate buffer and suspended in 1.5% Agarose, Low Melting Point, Analytical Grade (Promega, Madison, WI). After cooling at 4°C for 30 min, the agarose blocks were cut with a razor blade into thin slices. The slices were then stained with 1% KMnO4 at room temperature for an hour. After staining, slices were washed in distilled H2O, treated with 0.5% sodium meta-periodate for 15 min at room temperature, and washed again in distilled H2O. The slices were then dehydrated with a series of ethanol solutions (30, 50, 70, 95, and 100%) for 10 min each and left in 100% ethanol overnight. The next day, slices were washed in 100% ethanol and treated twice with propylene oxide for 10 min each time. Slices were then treated with differing ratios of propylene oxide to Spurr’s resin (Spurr, 1969). First, 3:1 for 3 h, followed by 1:1 overnight, 1:3 for 3 h, and finally 100% Spurr’s resin for 3 h, three times each. The slices were vacuum infiltrated overnight and then embedded in silicone molds and polymerized in an oven at 60°C for 24 h. Eighty nanometer ultrathin sections from the resin blocks were cut with a Reichert-Jung UltracutE ultramicrotome and placed on Formvar-coated slot copper grids. A Tecnai12 BioTwinG2 electron microscope (FEI, Hillsboro, OR), and an XR-60 CCD Digital Camera System (Advanced Microscopy Techniques, Woburn, MA) were used for capturing TEM images. Sample sectioning and TEM analyses were carried out at the Central Microscopy Imaging Center at Stony Brook University.

Assays for growth inhibition and invasive growth into agar

Spot assays were employed to assess the sensitivity of cells to high temperature (42°C), Calcofluor White, Congo Red, and SDS. Cells were grown overnight at 30°C to log phase and diluted to 1.0 × 107 cells/ml, which is the starting concentration of the 10-fold dilution series. With a multichannel pipette, 3 μl of each dilution was spotted on YPD plates containing 35 μg/ml Calcofluor White, 35 μg/ml Congo Red, or 100 μg/ml SDS. The agar plates were incubated at either 37 or 42°C and photographed after 48 h of growth.

To test the susceptibility of mutant strains to the PM stress agents duramycin and copper, disk diffusion assays were utilized. Cells were grown overnight to log phase in synthetic medium at 30°C, diluted to 1.0 × 107 cells/ml and 250 μl of the dilution was spread onto synthetic medium agar plate. Paper filter disks (Becton Dickinson and Company, Sparks, MD) were saturated with 10 μl of the indicated concentration of duramycin (Sigma Aldrich, St. Louis, MO) or CuSO4 and placed onto the surface of agar plates containing a lawn of the appropriate cell type. The plates were incubated for 48 h at 30°C and then the diameter of the zone of growth inhibition around the disk was recorded. Statistical tests were performed, and graphs were generated using Prism 6 software (GraphPad Software, La Jolla, CA.)

To measure invasive growth into agar, cells were grown overnight to log phase in synthetic medium at 30°C and diluted to 1.0 × 107 cells/ml. Three microliters of the dilution was spotted onto a petri plate containing 1.5% agar and 4% bovine serum. The plates were incubated at 37°C for 1 wk and then photographed (Naseem et al., 2020).

Supplementary Material

ACKNOWLEDGMENTS

We thank the members of our lab for their helpful advice and suggestions on the manuscript. This research was supported by a Public Health Service grant awarded to J.B.K. from the National Institutes of Health (R01AI047837). C.E.L. was supported in part by a National Institutes of Health training grant (NIH T32AI007539) from the National Institute of Allergy and Infectious Diseases.

Abbreviations used:

ANOVA analysis of variance

BAR Bin/amphiphysin/Rvs

GFP green fluorescent protein

GlcNAc N-Acetylglucosamine

MCC membrane compartment of Can1

PH pleckstrin homology

PI 4,5P 2 phosphatidylinositol 4,5-bisphosphate

PLC phospholipase C

PM plasma membrane

RFP red fluorescent protein

SDS sodium dodecyl sulfate

YPD yeast extract peptone dextrose

Reviewer Report

This article was published online ahead of print in MBoC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E23-08-0324) on May 22, 2024.
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