*Article* **Defining Functions of Mannoproteins in** *Saccharomyces cerevisiae* **by High-Dimensional Morphological Phenotyping**

**Farzan Ghanegolmohammadi 1,2 , Hiroki Okada <sup>3</sup> , Yaxuan Liu <sup>1</sup> , Kaori Itto-Nakama <sup>1</sup> , Shinsuke Ohnuki <sup>1</sup> , Anna Savchenko 1,4, Erfei Bi <sup>3</sup> , Satoshi Yoshida <sup>5</sup> and Yoshikazu Ohya 1,\***


**Abstract:** Mannoproteins are non-filamentous glycoproteins localized to the outermost layer of the yeast cell wall. The physiological roles of these structural components have not been completely elucidated due to the limited availability of appropriate tools. As the perturbation of mannoproteins may affect cell morphology, we investigated mannoprotein mutants in *Saccharomyces cerevisiae* via high-dimensional morphological phenotyping. The mannoprotein mutants were morphologically classified into seven groups using clustering analysis with Gaussian mixture modeling. The pleiotropic phenotypes of cluster I mutant cells (*ccw12*∆) indicated that *CCW12* plays major roles in cell wall organization. Cluster II (*ccw14*∆, *flo11*∆, *srl1*∆, and *tir3*∆) mutants exhibited altered mother cell size and shape. Mutants of cluster III and IV exhibited no or very small morphological defects. Cluster V (*dse2*∆, *egt2*∆, and *sun4*∆) consisted of endoglucanase mutants with cell separation defects due to incomplete septum digestion. The cluster VI mutant cells (*ecm33*∆) exhibited perturbation of apical bud growth. Cluster VII mutant cells (*sag1*∆) exhibited differences in cell size and actin organization. Biochemical assays further confirmed the observed morphological defects. Further investigations based on various omics data indicated that morphological phenotyping is a complementary tool that can help with gaining a deeper understanding of the functions of mannoproteins.

**Keywords:** mannoprotein; cell wall; budding yeast; morphology; CalMorph

## **1. Introduction**

The cell wall is a rigid structure that plays essential roles in establishing cell morphology and dictating the oval shape of budding yeast, *Saccharomyces cerevisiae*, and it also confers robustness on the cell by stabilizing internal osmotic conditions and serving as a site for cell wall enzymes to exert their effects [1–3]. Electron microscopic analysis has revealed that the yeast cell wall is a highly organized composite consisting of internal interconnected filamentous polysaccharides (1,3-β-glucan, 1,6-β-glucan, and chitin) and external non-filamentous glycoproteins (mannoproteins), which form a firm extracellular matrix similar to reinforced concrete [4,5]. Whereas 1,3-β-glucan is the major filamentous cell wall component [2,6] mainly responsible for dictating the yeast cell shape, non-filamentous mannoproteins, of which 36 members have been identified to date, have also been suggested to play fundamental roles in the cell wall [3].

**Citation:** Ghanegolmohammadi, F.; Okada, H.; Liu, Y.; Itto-Nakama, K.; Ohnuki, S.; Savchenko, A.; Bi, E.; Yoshida, S.; Ohya, Y. Defining Functions of Mannoproteins in *Saccharomyces cerevisiae* by High-Dimensional Morphological Phenotyping. *J. Fungi* **2021**, *7*, 769. https://doi.org/10.3390/jof7090769

Academic Editors: María Molina and Humberto Martín

Received: 28 July 2021 Accepted: 14 September 2021 Published: 17 September 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

Previous studies have indicated that individual deletions of genes encoding mannoproteins may result in subtle growth defects [7–10]. This can be partly explained by gene duplication, as 26 of 36 mannoprotein genes are duplicated [11]. Another reason is that cell wall defects caused by the lack of mannoproteins affect cell morphology rather than growth phenotypes [12]. Thus, the morphological phenotyping of mannoprotein mutants would provide more information on their functions, highlighting the importance of morphology as another metric with which to study the genes involved in cell wall assembly. In general, mannoproteins play a collective role in maintaining the cell wall structure [13], but differences in the localization, structure, and probably also the function of mannoproteins in the cell wall [3,8] suggest that the perturbation of individual genes may result in different morphological phenotypes [12]. Each mannoprotein is likely to have a distinct role in the cell wall, but the details have not been elucidated due to limited quantitative morphological analysis of mannoprotein mutants.

This study was performed to determine a responsibility assignment matrix (hereafter we call it responsibility matrix) through morphological clustering analysis of mannoproteins in relation to their molecular functions. For this purpose, high-dimensional morphological phenotyping was performed after extracting the morphological features of each mannoprotein mutant with the image processing program CalMorph [14]. Analyses of morphological abnormalities based on a powerful parametric approach revealed specific morphological phenotypes that will help with uncovering the responsibility matrix of mannoproteins in the yeast cell wall.

#### **2. Materials and Methods**

## *2.1. Strains and Growth Conditions*

Of 36 mannoprotein gene-deletion mutants of the budding yeast, *S. cerevisiae* [3], we studied 32 mutants (Table S1) that were straightforward to observe as single cells under a microscope. These 32 mutants were isogenic derivatives of BY4741 (*MATa his3 leu2 met15 ura3*) and were purchased from EUROSCARF (Oberursel, Germany). Cells with mutations in the other four mannoprotein genes (*aga1*∆, *flo5*∆, *flo9*∆, and *dan4*∆) were not studied due to heavy cell aggregation. Mutant and wild-type (WT) strains were cultured under optimal growth conditions at 25 ◦C in nutrient-rich yeast extract peptone dextrose (YPD) medium containing 1% (*w*/*v*) Bacto yeast extract (BD Biosciences, San Jose, CA, USA), 2% (*w*/*v*) polypeptone (Wako Chemicals, Richmond, VA, USA), and 2% (*w*/*v*) dextrose, as described previously [14]. A WT diploid strain (BY4743) and the homozygous gene deletion mutants in the BY4743 background used for Western blotting were purchased from EUROSCARF (Oberursel, Germany).

#### *2.2. Fluorescence Staining, Microscopy, and Image Processing*

To minimize variation due to inconsistencies in data acquisition, we followed a precise protocol for the preparation, fixation, and fluorescence staining of yeast cells in the early *log* phase of growth (<5.0 <sup>×</sup> <sup>10</sup><sup>6</sup> cells; 5 biological replicates), as described previously [15–18]. Briefly, yeast cells were fixed for 30 min in growth medium supplemented with formaldehyde (final concentration, 3.7%) and potassium phosphate buffer (100 mM, pH 6.5) at 25 ◦C. Yeast cells were then collected via centrifugation at room temperature and further incubated in potassium phosphate buffer containing 4% formaldehyde for 45 min. The fixed cells were subsequently prepared for fluorescence microscopy. First, actin staining was performed by treating the cells overnight with 15 U/mL rhodaminephalloidin (Invitrogen, Carlsbad, CA, USA) and 1% Triton-X in phosphate-buffered saline (PBS). Second, cell wall mannoproteins were stained by treating cells for 10 min with 1 mg/mL fluorescein isothiocyanate-conjugated concanavalin A (Sigma-Aldrich, St. Louis, MO, USA) in P buffer (10 mM sodium phosphate and 150 mM NaCl, pH 7.2). Finally, after washing twice with P buffer, the yeast cells were mixed with mounting buffer (1 mg/mL *p*-phenylenediamine, 25 mM NaOH, 10% PBS, and 90% glycerol) containing 20 mg/mL 4 0 ,6-diamidino-2-phenylindole (Sigma-Aldrich) to stain DNA.

Images of triple-stained cells were captured using an Axio Imager microscope equipped with a 6100 ECplan-Neofluar lens (Carl Zeiss, Oberkochen, Germany), a CoolSNAP HQ cooled charged coupled device (CCD) camera (Roper Scientific Photometrics, Tucson, AZ, USA), and AxioVision software ver. 4.5 (Carl Zeiss). The obtained images were quantified using CalMorph with regard to 501 morphological parameters related to the cell-cycle phase, actin cytoskeleton, cell wall, and nuclear DNA. The descriptions for each trait have been reported previously [14], and the CalMorph user manual is available at http://www. yeast.ib.k.u-tokyo.ac.jp/CalMorph/download.php?path=CalMorph-manual.pdf, accessed on 21 September 2019. Only those experiments containing at least 200 cells, detected by CalMorph, were considered for statistical analysis.

#### *2.3. Data Analysis*

All statistical analyses were performed using R (http://www.r-project.org, accessed on 21 September 2019). To assess the effects of genetic perturbation on the morphology of the mutants, we compared the cell morphological traits of the mutants with the corresponding WT distribution (i.e., null distribution) for each trait using an ANOVA model based on a generalized linear model (GLM). The GLM is an extension of the normal linear model in which predictors are linear but link functions are nonlinear to cope with violations of some standard assumptions of linear models [19]. These properties allow the analysis to cover probability distributions other than the Gaussian distribution. CalMorph generated 501 morphological parameters with which we established models based on the probability distributions for 490 unimodal parameters using the UNImodal MOrphological data pipeline (UNIMO; unpublished). Briefly, we first categorized CalMorph parameters into the following four data types: non-negative parameters, ratios, coefficients of variation (CVs; further converted to noise values, see below), and proportions. Then, we showed that these parameters could be explained well by 10 unimodal distributions to accommodate the statistical model used in the GLM: gamma, inverse gamma, and Weibull distributions for non-negative parameters; beta and logit-normal distributions for ratios; Gaussian, logistic, and reverse Gumbel distributions for noise parameters; and binomial and beta-binomial distributions for proportions. The best fit unimodal probability distribution for each parameter was eventually determined using the Akaike information criterion (AIC). CVs ( *Population standard deviation* (*σ*) *Population mean* (*µ*) ) are nonlinearly dependent on mean values [9]. We used LOESS (locally estimated scatterplot smoothing) regression with a smooth span (*f*) to uncouple this concomitant dependency. AIC values were used to choose the best-fitting model among various smooth spans (0.10 ≤ *f* ≤ 0.99). Finally, noise parameters were calculated as the residuals, i.e., observed value minus predicted value.

To estimate *Z*-values, once maximum-likelihood estimation converged, we transformed each morphological parameter to a *Z*-value via the Wald test (one-sample two-sided test) using the summary.gamlss R function [20]. The false discovery rate (FDR), the rate of type I error associated with rejecting the null hypothesis due to multiple comparisons, was estimated based on 2000 permutations.

#### *2.4. Dimensionality Reduction and Clustering*

To extract the most effective parameters, we performed principal component analysis (PCA), the most commonly used method for reducing dimensionality [21,22], on the obtained *Z*-values using the prcomp function (stats package). We then calculated the cumulative contribution ratio (CCR) to describe variation in the data. Based on the result, we used the first five principal components (CCR = 81.34%) for clustering analysis (Figure S1A).

Mixture model clustering is a probability-based approach in which we assume the dataset is best described as a mixture of probability models. In Gaussian mixture modeling (GMM), the most commonly used model-based clustering method [23], Gaussian distributions are fitted to the dataset. Gaussian distributions are randomly initialized and their parameters optimized iteratively to achieve a better fit. The expectation maximization algo-

rithm estimates all parameters to assign members into *c* clusters. We employed the mclust package [23] to determine the underlying Gaussian mixture distributions (Figure S1B,C).

#### *2.5. Kinetics of Cluster V Mannoproteins (Dse2 and Egt2)*

#### 2.5.1. Yeast Media and Culture Conditions

Standard culture media and genetic techniques were used [24]. Yeast strains were grown routinely at 25 ◦C in synthetic complete (SC) minimal medium lacking specific amino acid(s) and/or uracil or YPD. Neutralized SC medium (pH 7.0) was used for livecell imaging of green fluorescent protein (GFP) molecules exposed to the extracellular environment to prevent quenching of the GFP signal caused by the acidity of the standard SC medium.

#### 2.5.2. Constructions of Strains

New strains were constructed either by integrating a plasmid carrying a modified gene at a genomic locus or by transferring a deletion or tagged allele of a gene from a plasmid or from one strain to another via PCR amplification and yeast transformation; see footnotes in Table S2 [25–27].

#### 2.5.3. Primers and Plasmids

All PCR primers and plasmids used in this study are listed in Table S3. All PCR primers were purchased from Integrated DNA Technologies (Coralville, IA, USA). All new constructs were validated via sequencing performed at the DNA Sequencing Facility, University of Pennsylvania. The plasmids pFA6a-GFPEnvy-KanMX6, pFA6a-link-GFPEnvy-KanMX6, and pRS316-ENVY-FKS1(1-789) were described previously [28]. The plasmids bWL715 (pHIS3p:mRuby2-Tub1+3'UTR::HPH [29]) and pFA6a-URA3-KanMX6 [30] were generous gifts from Wei-Lih Lee (Dartmouth College) and John Pringle (Stanford University), respectively.

The following plasmids were generated for this study. To generate pFA6a-link-GFPEnvy-CaURA3, a ~0.7-kb *Pac*I-*Asc*I fragment containing *GFPEnvy* from pFA6a-link-GFPEnvy-SpHis5 [27] was subcloned to replace the ~0.7-kb *Pac*I-*Asc*I region of pFA6alink-yomApple-CaURA3 (#44879; Addgene, Watertown, MA, USA). To generate proHIS3 ymScarlet-I-TUB1-tTUB1-HPH (integrative, *hphMX*, expresses Tub1 N-terminally tagged with ymScarlet-I under the control of the *HIS3* promoter), two DNA fragments carrying either the ~0.7-kb ymScarlet-I insert or a ~6.5-kb plasmid backbone were amplified via PCR using the plasmid YIp128-proACT1-lifeact-ymScarlet-I-tADH1 (lab stock, integrative, *LEU2*, expresses Lifeact C-terminally tagged with ymScarlet-I under the control of the *ACT1* promoter) as the template DNA and the primers P1409 and P1412, or using the plasmid bWL715 as the template DNA and the primers P1410 and P1411, respectively. The resultant PCR products were then assembled using a Quick-Fusion cloning kit (Bimake, Houston, TX, USA). To generate pRS305-ENVY-FKS1(1-789), a ~4.2-kb DNA fragment carrying the partial open reading frame (ORF) of *GFPEnvy-FKS1* (from ~1 kb of the *FKS1* promoter region, *GFPEnvy*, and the *FKS1* ORF until residue 789 followed by a new stop codon) was amplified via PCR using pRS316-ENVY-FKS1(1-789) as the template DNA and the primers P222 and P512. The resultant PCR product was then subcloned into *Apa*I- and *Sac*I-digested pRS305 (integrative, *LEU2*) using a Quick-Fusion cloning kit.

#### 2.5.4. Imaging and Data Analysis

Time-lapse microscopy was conducted as described previously with slight modifications [31]. Cells were cultured to an exponential phase at 25 ◦C in SC medium, briefly sonicated at 15% power for 5 s to declump the cells (model Q55; Qsonica, Newtown, CT, USA), concentrated via centrifugation, and spotted onto concanavalin A-coated glassbottom dishes. After a sufficient amount of cells had adhered to the bottom of each dish (> 50% cell cover in a microscopic field), the SC medium was replaced with neutralized SC liquid medium, and the dishes were then incubated at room temperature (23 ◦C) for

15 min to allow the cells to acclimatize. Images were acquired at room temperature with a spinning-disk confocal microscope (Eclipse Ti2-U; Nikon, Tokyo, Japan) with a 100× /1.49NA oil objective (CFI Apo TIRF 100×; Nikon) combined with a confocal scanner unit (CSU-X1; Yokogawa, Tokyo, Japan). An EMCCD camera (Evolve 512 Delta; Photometrics, Tucson, AZ, USA) was used for image capturing. Solid-state lasers for excitation (488 nm for GFP, and 561 nm for red fluorescent protein) were housed in a laser merge module (ILE-400; Spectral Applied Research, Richmond Hill, ON, Canada). The imaging system was controlled using MetaMorph (version 7.10.4.431, Molecular Devices, San Jose, CA, USA). Images were taken every 2 min with 11 z-stacks with a step size of 0.8 µm. Sum or maximum intensity projections were calculated using NIH ImageJ (1.51 h) [32]. To quantify fluorescence intensities, the integrated density at a division site was calculated from the sum intensity projection of an image stack by subtracting the fluorescence intensity in the background area from the total intensity in an ImageJ-drawn polygon covering the division site.

#### *2.6. Biochemistry*

Whole-cell protein extracts were prepared as described previously [33]. Briefly, cells were pelleted, treated with NaOH (0.1 N), and incubated on ice (5 min). Then, cells were pelleted, resuspended in SDS sample buffer including 62.5 mM Tris-HCl (pH 6.8), glycerol (10%), SDS (2%), β-mercaptoethanol (2%), and bromophenol blue (0.005%), boiled for 5 min, and pelleted. Afterward, the supernatants were loaded in a mini-gel (4–15%; Bio-Rad, CA, USA), and Western blotting was performed with rabbit anti–phospho-p42/44 MAPK (T202/Y204) antibody (Cell Signaling Technology, Danvers, MA, USA) and rabbit anti-yeast Rho1 antibody (Abmart, Berkeley Heights, NJ, USA). HRP-conjugated secondary antibodies were obtained from Millipore, and proteins were detected with an enhanced chemiluminescence system (ECL plus; Amersham, Darmstadt, Germany).

#### *2.7. Similarity of Mannoprotein Mutants and Drug-Treated Wild-Type Cells in Morphology*

Morphological profiles of ccw12∆ (I), ccw14∆ (II), cwp2∆ (IV), sun4∆ (V), ecm33∆ (VI), and sag1∆ (VII) were compared with WT cells treated with unicamycin, echinocandin B, nikkomycin Z, and hydroxyurea. Morphological data of the drug-treated cells were obtained from [34]. To investigate the profile similarity, first, CalMorph values were transformed to *Z*-values (Wald test) using the UNIMO pipeline (490 parameters). Then, the obtained *Z*-values of the WT replicates were exposed to PCA. Finally, *Z*-values of the mutants/drug-treated cells were projected onto PC axes of the WT. Pearson correlation coefficient (r) was calculated between each pair using first 94 PC scores (CCR = 99%).

#### *2.8. Mannoprotein Analysis Based on Omics Studies*

#### 2.8.1. Estimation of Fitness

To estimate the fitness of 32 mannoprotein mutants, a previously reported dataset containing the logarithmic strain growth rate coefficients of gene-deletion mutants grown on basal medium (LSCbasal) was employed [10]. *p*-values were calculated to determine whether the fitness of each strain was significantly lower than that of the WT based on one tail of the estimated probability distribution, as described previously [12], using the pnorm function (stats package), and the results were corrected for family-wise error using the qvalue function in the qvalue package [35].

#### 2.8.2. Analysis of Protein Abundance and Protein–Protein Interactions

To determine the abundance of 32 mannoproteins at the protein level, mean values from 21 datasets were used as reported previously [36]. Protein–protein interaction (PPI) data were obtained from the BioGRID database [37]. We examined physical interactions (between interactors A and B) for each mutant of *S. cerevisiae* S288C (Taxonomy ID: 559292). Two types of PPI networks were considered: PPIs among the 32 mannoproteins and PPIs

between each of the 32 mannoproteins and the whole proteome (i.e., the protein–protein interactome profile). Networks were visualized using Cytoscape 3.8.2 [38].

#### 2.8.3. Genetic Interaction Analysis

Genetic interaction (GI) data were collected as reported previously [39]. Significant interactions based on both queries and array analysis were considered for further analysis (*p* < 0.05). Two types of GI networks were considered: GIs among the 32 mannoprotein genes and GIs between each of the 32 mannoprotein genes and the whole genome (i.e., the genetic interactome profile). Finally, networks were visualized using Cytoscape 3.8.2 [38].

#### 2.8.4. Chemical-Genetic Profile Analysis

The chemical-genetic profiles of the 32 mannoproteins were obtained through text mining of the *Saccharomyces* Genome Database (SGD).

#### **3. Results**

#### *3.1. Effects of Genetic Perturbations on Cell Morphology*

We analyzed the morphology of mutants with deletions of individual genes encoding 32 mannoproteins using the image processing program CalMorph. To perform morphological phenotyping, it is necessary to consider the diversity in yeast morphological measurements. We applied different probability distribution models to accurately estimate the true value of each morphological parameter [17]. The use of 490 unimodal morphological parameters enabled a powerful approach, revealing biological information that may be masked with commonly used imaging methods. We found that cell morphology was remarkably altered: of 490 parameters, perturbations were detected in 136 parameters, consisting of 16, 77, and 43 parameters related to actin, cell, and nuclear DNA morphology, respectively (Wald test, FDR = 0.05; Table S4). This observation implies profound effects of mannoproteins on cell morphology, suggesting that mannoproteins may play roles in dictating cell shape and the progression of the cell cycle.

To understand the morphological alterations more holistically, we reduced the number of dimensions of the morphological space to five via PCA of the Z-values of 136 significantly changed parameters; the first five principal components accounted for 81.34% of the variation (Figure S1A and Table S5). We then used GMM, one of the most commonly used model-based clustering methods for normally distributed data, to cluster the mannoprotein mutants (Figure S1B). The posterior probabilities associated with the data were evaluated in our GMM analysis to validate our clustering results (Figure S1C). Using GMM, we successfully clustered 32 mannoprotein mutants into seven groups (Figure 1). The *ccw12*∆ mutant, the single member of cluster I, was the mutant with the most abundant covalently linked cell wall protein. Members of cluster II (*ccw14*∆, *flo11*∆, *srl1*∆, and *tir3*∆) were mutants of serine-rich mannoproteins. Cluster III (nine mutants) and IV (13 mutants) accounted for more than half of the mannoprotein mutants, with their members exhibiting no or very small effects on cell morphology. Members of cluster V (*dse2*∆, *egt2*∆, and *sun4*∆) were endoglucanase mutants. The *ecm33*∆ mutant in cluster VI had a mutation in a glycosylphosphatidylinositol (GPI)-anchored protein thought to be involved in bud morphogenesis. In the single member of cluster VII, the *sag1*∆ mutant, no morphological defects in vegetative growth had been reported previously.

bly.

**Figure 1.** Two-dimensional principal component (PC) analysis score plot (biplot) illustrating clustering of the 32 mannoproteins. Each circle represents a single mannoprotein deletion mutant. The mixture likelihood values at individual points, based on the first five PC scores (CCR = 81.34%; Figure S1A) and a seven-component EEI model (Figure S1B), revealed data trends, including seven **Figure 1.** Two-dimensional principal component (PC) analysis score plot (biplot) illustrating clustering of the 32 mannoproteins. Each circle represents a single mannoprotein deletion mutant. The mixture likelihood values at individual points, based on the first five PC scores (CCR = 81.34%; Figure S1A) and a seven-component EEI model (Figure S1B), revealed data trends, including seven clusters. Mutants are color-coded.

#### clusters. Mutants are color-coded. *3.2. Phenotype of the Cluster I Mutant (ccw12*∆*)*

*3.2. Phenotype of the Cluster I Mutant (ccw12*Δ*)* Among all mannoprotein mutants, *ccw12*Δ cells exhibited the greatest morphological alterations with 81 significantly changed parameters (Wald test, FDR = 0.05; Table S6). The *ccw12*Δ cells were larger in size at the S/G2 (C11-1\_A1B and C101\_A1B) and M phases (C11-1\_C) and had a rounder cell shape (C115\_A, C115\_A1B, and C115\_C) and wider neck at both the S/G2 (C109\_A1B) and M phases (C109\_C) (Figure S2). In addition, the *ccw12*Δ mutation affected bud morphogenesis, resulting in a rounded bud shape (C114\_A1B and C114\_C) and a disturbed budding direction (C106\_A1B and C106\_C) (Figure S3). Further Among all mannoprotein mutants, *ccw12*∆ cells exhibited the greatest morphological alterations with 81 significantly changed parameters (Wald test, FDR = 0.05; Table S6). The *ccw12*∆ cells were larger in size at the S/G2 (C11-1\_A1B and C101\_A1B) and M phases (C11- 1\_C) and had a rounder cell shape (C115\_A, C115\_A1B, and C115\_C) and wider neck at both the S/G2 (C109\_A1B) and M phases (C109\_C) (Figure S2). In addition, the *ccw12*∆ mutation affected bud morphogenesis, resulting in a rounded bud shape (C114\_A1B and C114\_C) and a disturbed budding direction (C106\_A1B and C106\_C) (Figure S3). Further phenotypic analysis using chitin staining revealed a significantly elevated population of cells exhibiting abnormal chitin staining (*p* < 0.05, *t*-test, Figure 2A,B), demonstrating that the loss of *CCW12* function had a detrimental impact on cell wall organization and assembly.

phenotypic analysis using chitin staining revealed a significantly elevated population of cells exhibiting abnormal chitin staining (*p* < 0.05, *t*-test, Figure 2A,B), demonstrating that the loss of *CCW12* function had a detrimental impact on cell wall organization and assem-Cell wall damage is accompanied by the activation of the cell wall integrity (CWI) pathway and the phosphorylation of Slt2 MAPK [40–42]. We found that the *ccw12*∆ mutant exhibited a marked increase in Slt2 phosphorylation, indicating that the cell wall was damaged in this mutant (Figures 3 and S4).

Cell wall damage is accompanied by the activation of the cell wall integrity (CWI)

pathway and the phosphorylation of Slt2 MAPK [40–42]. We found that the *ccw12*Δ mutant exhibited a marked increase in Slt2 phosphorylation, indicating that the cell wall was

Taken together, and given that Ccw12 is important for CWI, these observations indicate that this gene deletion causes pleiotropic defects in cell growth and morphology, possibly because of a severe loss of mannoprotein structures and functions. Taken together, and given that Ccw12 is important for CWI, these observations indicate that this gene deletion causes pleiotropic defects in cell growth and morphology, possibly because of a severe loss of mannoprotein structures and functions.

*J. Fungi* **2021**, *7*, x FOR PEER REVIEW 8 of 19

**Figure 2.** Abnormalities in *ccw12*Δ (cluster I) and *ecm33*Δ (cluster VI) cells. (**A**). Wild-type (WT) and mutant cells were grown in yeast extract peptone dextrose medium at 25 °C with shaking at 200 rpm until *log* phase. Cells (2.0 × 10<sup>6</sup> cells) were suspended in 1 mL of phosphate-buffered saline (PBS) and mixed well with 5 μL of 5 mg/mL wheat germ agglutinin in PBS to stain chitin. After incubation at room temperature (30 min), the stained cells were washed three times and observed under a fluorescence microscope with a 4′,6-diamidino-2-phenylindole filter. The bar plot shows the percentages of abnormal *ccw12*Δ (cluster I) and *ecm33*Δ (cluster VI) cells in comparison with WT cells. Error bars indicate standard deviations. \* *p* < 0.05 (*t* test). (**B**). Examples of chitin staining in **Figure 2.** Abnormalities in *ccw12*∆ (cluster I) and *ecm33*∆ (cluster VI) cells. (**A**). Wild-type (WT) and mutant cells were grown in yeast extract peptone dextrose medium at 25 ◦C with shaking at 200 rpm until *log* phase. Cells (2.0 <sup>×</sup> <sup>10</sup><sup>6</sup> cells) were suspended in 1 mL of phosphate-buffered saline (PBS) and mixed well with 5 µL of 5 mg/mL wheat germ agglutinin in PBS to stain chitin. After incubation at room temperature (30 min), the stained cells were washed three times and observed under a fluorescence microscope with a 40 ,6-diamidino-2-phenylindole filter. The bar plot shows the percentages of abnormal *ccw12*∆ (cluster I) and *ecm33*∆ (cluster VI) cells in comparison with WT cells. Error bars indicate standard deviations. \* *p* < 0.05 (*t* test). (**B**). Examples of chitin staining in WT, *ccw12*∆ (cluster I), and *ecm33*∆ (cluster VI) cells.

WT, *ccw12*Δ (cluster I), and *ecm33*Δ (cluster VI) cells.

*J. Fungi* **2021**, *7*, x FOR PEER REVIEW 9 of 19

model clustering of morphological data (see Figure 1).

age (Figure 3 and Figure S4).

**Figure 3.** Western blotting of phosphorylated Slt2 (pSlt2, upper panel) and loading control Rho1 (lower panel). BY4743 (WT); *ccw12*Δ/*ccw12*Δ (cluster I); *ccw14*Δ/*ccw14*Δ, *flo11*Δ/*flo11*Δ, *srl1*Δ/*srl1*Δ, and *tir3*Δ/*tir3*Δ (cluster II); *hsp150*Δ/*hsp150*Δ (cluster IV); *dse2*Δ/*dse2*Δ, *egt2*Δ/*egt2*Δ, and *sun4*Δ/*sun4*Δ (cluster V); *ecm33*Δ/*ecm33*Δ (cluster VI); and *sag1*Δ/*sag1*Δ (cluster VII) cells were examined for the presence of phosphorylated Slt2. Rabbit antibody against phospho-p42/44 MAPK (T202/Y204) and rabbit antibody against yeast Rho1 were used to detect the phosphorylated Slt2 and Rho1, respectively. *slt2*Δ/*slt2*Δ and *sac7*Δ/*sac7*Δ were used as negative and positive controls, respectively, for phosphorylated Slt2. Mutants are color-coded according to Gaussian mixture **Figure 3.** Western blotting of phosphorylated Slt2 (pSlt2, upper panel) and loading control Rho1 (lower panel). BY4743 (WT); *ccw12*∆/*ccw12*∆ (cluster I); *ccw14*∆/*ccw14*∆, *flo11*∆/*flo11*∆, *srl1*∆/*srl1*∆, and *tir3*∆/*tir3*∆ (cluster II); *hsp150*∆/*hsp150*∆ (cluster IV); *dse2*∆/*dse2*∆, *egt2*∆/*egt2*∆, and *sun4*∆/*sun4*∆ (cluster V); *ecm33*∆/*ecm33*∆ (cluster VI); and *sag1*∆/*sag1*∆ (cluster VII) cells were examined for the presence of phosphorylated Slt2. Rabbit antibody against phospho-p42/44 MAPK (T202/Y204) and rabbit antibody against yeast Rho1 were used to detect the phosphorylated Slt2 and Rho1, respectively. *slt2*∆/*slt2*∆ and *sac7*∆/*sac7*∆ were used as negative and positive controls, respectively, for phosphorylated Slt2. Mutants are color-coded according to Gaussian mixture model clustering of morphological data (see Figure 1).

## *3.3. Phenotype of Cluster II Mutants (ccw14*∆*, srl1*∆*, flo11*∆*, and tir3*∆*)*

*3.3. Phenotype of Cluster II Mutants (ccw14*Δ*, srl1*Δ*, flo11*Δ*, and tir3*Δ*)* Cluster II mutants tended to produce larger mother cells at the M phase. The most noticeable morphological mutant in this cluster was *srl1*∆, which had a significantly larger mother cell size (C11-1\_C), mother cell outline length (C12-1\_C), and long axis (C103\_C; Wald test, FDR = 0.05; Table S6). Both the mother cell outline length (C12-1\_C) and long axis length (C103\_C) of all cluster II mutants were larger than those in the other clusters, and nearly equivalent to those of the cluster I mutant (*ccw12*∆) (Figure S5). Therefore, we considered that the cluster II mutants exhibited perturbations in the mother cell size and shape at the M phase. There was no obvious increase in Slt2 phosphorylation, suggesting Cluster II mutants tended to produce larger mother cells at the M phase. The most noticeable morphological mutant in this cluster was *srl1*∆, which had a significantly larger mother cell size (C11-1\_C), mother cell outline length (C12-1\_C), and long axis (C103\_C; Wald test, FDR = 0.05; Table S6). Both the mother cell outline length (C12-1\_C) and long axis length (C103\_C) of all cluster II mutants were larger than those in the other clusters, and nearly equivalent to those of the cluster I mutant (*ccw12*∆) (Figure S5). Therefore, we considered that the cluster II mutants exhibited perturbations in the mother cell size and shape at the M phase. There was no obvious increase in Slt2 phosphorylation, suggesting little cell wall damage in the cluster II mutants (Figures 3 and S4).

#### little cell wall damage in the cluster II mutants (Figure 3 and Figure S4). *3.4. Phenotype of Cluster V Mutants (dse2*∆*, egt2*∆*, and sun4*∆*)*

*3.4. Phenotype of Cluster V Mutants (dse2*∆*, egt2*∆*, and sun4*∆*)* Among the cluster V mutants, *egt2*∆ exhibited the greatest morphological changes, with significant differences in 31 parameters (Wald test, FDR = 0.05, Table S6). Morphological analysis of the cluster V mutants revealed common morphological features, such as the accumulation of cells at the M phase (D202 and D213) with actin patches localized at the bud neck (A109 and A118) (Figure S6). As actin patches are localized to the bud neck in cytokinesis, the morphological features of the cluster V mutants are suggestive of defects in cell separation. Consistent with this, cluster V genes (*DSE2, EGT2,* and *SUN4*) all encode cell wall mannoproteins similar to glucanase. It should be noted that the mutants exhibited no significant changes in bud cell size (C11-2\_C and C12-2\_C) or nuclear size (D14-2\_C and D17-2\_C) (Figure S7A), suggesting no defects in cell division but defects in physical attachment between mother and daughter cells. Mother cells frequently started the next budding cycle while still attached to old daughter cells (Figure S7B). The phosphorylation of Slt2 was increased in all cluster V mutants, suggesting cell wall dam-Among the cluster V mutants, *egt2*∆ exhibited the greatest morphological changes, with significant differences in 31 parameters (Wald test, FDR = 0.05, Table S6). Morphological analysis of the cluster V mutants revealed common morphological features, such as the accumulation of cells at the M phase (D202 and D213) with actin patches localized at the bud neck (A109 and A118) (Figure S6). As actin patches are localized to the bud neck in cytokinesis, the morphological features of the cluster V mutants are suggestive of defects in cell separation. Consistent with this, cluster V genes (*DSE2, EGT2,* and *SUN4*) all encode cell wall mannoproteins similar to glucanase. It should be noted that the mutants exhibited no significant changes in bud cell size (C11-2\_C and C12-2\_C) or nuclear size (D14-2\_C and D17-2\_C) (Figure S7A), suggesting no defects in cell division but defects in physical attachment between mother and daughter cells. Mother cells frequently started the next budding cycle while still attached to old daughter cells (Figure S7B). The phosphorylation of Slt2 was increased in all cluster V mutants, suggesting cell wall damage (Figures 3 and S4).

Glucanases are localized at the site of division in cytokinesis. To understand the precise timing of the function of glucanases in cell separation, we tagged cluster V gene products with GFP and performed quantitative time-lapse imaging to obtain information on

Glucanases are localized at the site of division in cytokinesis. To understand the precise timing of the function of glucanases in cell separation, we tagged cluster V gene products with GFP and performed quantitative time-lapse imaging to obtain information on real-time protein abundance at the division site (Figure 4) [28]. The accumulation peaks of both GFP-Egt2 and Dse2-GFP occurred after those of two secondary septum (SS)-forming enzymes, GFP-Fks1 and Chs3-GFP, suggesting that cluster V genes likely function after SS formation. Dse2-GFP exhibited accumulation kinetics remarkably similar to those of Cts1-GFP (*r* = 0.98, Table S7), a chitinase required for the degradation of the primary septum (PS) during cell separation. These observations imply that Dse2 may function in the same process as Cts1. Interestingly, the peak of GFP-Egt2 at the division site occurred between the peaks of the SS-forming enzymes and the peak of the PS-degrading enzyme, suggesting that Egt2 may be involved in cell wall remodeling or maturation, which is required for cell separation. Taken together, these results further support the involvement of cluster V genes in cell separation and explain the major cluster V mutant phenotype of mother cells with unseparated old daughter cells. *J. Fungi* **2021**, *7*, x FOR PEER REVIEW 10 of 19 real-time protein abundance at the division site (Figure 4) [28]. The accumulation peaks of both GFP-Egt2 and Dse2-GFP occurred after those of two secondary septum (SS)-forming enzymes, GFP-Fks1 and Chs3-GFP, suggesting that cluster V genes likely function after SS formation. Dse2-GFP exhibited accumulation kinetics remarkably similar to those of Cts1-GFP (*r* = 0.98, Table S7), a chitinase required for the degradation of the primary septum (PS) during cell separation. These observations imply that Dse2 may function in the same process as Cts1. Interestingly, the peak of GFP-Egt2 at the division site occurred between the peaks of the SS-forming enzymes and the peak of the PS-degrading enzyme, suggesting that Egt2 may be involved in cell wall remodeling or maturation, which is required for cell separation. Taken together, these results further support the involvement of cluster V genes in cell separation and explain the major cluster V mutant phenotype of mother cells with unseparated old daughter cells.

**Figure 4.** Kinetics of Dse2 and Egt2 (cluster V) proteins and proteins involved in cytokinesis and cell separation**.** (**A**). Images of green fluorescent protein (GFP)-tagged cluster V proteins and proteins involved in cytokinesis and cell separation. Montages of cells were created from frames selected from time-lapse series consisting of images taken at 2-min intervals. The white dotted line represents the cell outline. The following strains were used: YEF10861 (*MYO1-GFP mScarlet-TUB1*), YEF10856 (*CHS3-GFP mScarlet-TUB1*), YEF10857 (*GFP-FKS1 mScarlet-TUB1*), YEF10862 (*CTS1-GFP mScarlet-TUB1*), YEF10879 (*GFP-EGT2 mScarlet-TUB1*), and YEF10858 (*DSE2-GFP mScarlet-TUB1*). (**B**). Kinetics of the GFP-tagged proteins indicated in "**A**". The vertical dashed line shows timing of spindle breakage. Bold lines and associated shaded bands represent mean and SD values, respectively. *n* > 23 for each strain. *3.5. Phenotype of the Cluster VI Mutant (ecm33*∆*)* **Figure 4.** Kinetics of Dse2 and Egt2 (cluster V) proteins and proteins involved in cytokinesis and cell separation. (**A**). Images of green fluorescent protein (GFP)-tagged cluster V proteins and proteins involved in cytokinesis and cell separation. Montages of cells were created from frames selected from time-lapse series consisting of images taken at 2-min intervals. The white dotted line represents the cell outline. The following strains were used: YEF10861 (*MYO1-GFP mScarlet-TUB1*), YEF10856 (*CHS3-GFP mScarlet-TUB1*), YEF10857 (*GFP-FKS1 mScarlet-TUB1*), YEF10862 (*CTS1-GFP mScarlet-TUB1*), YEF10879 (*GFP-EGT2 mScarlet-TUB1*), and YEF10858 (*DSE2-GFP mScarlet-TUB1*). (**B**). Kinetics of the GFP-tagged proteins indicated in (**A**). The vertical dashed line shows timing of spindle breakage. Bold lines and associated shaded bands represent mean and SD values, respectively. *n* > 23 for each strain.

#### The *ecm33*∆ cells exhibited significant differences in 22 morphological parameters *3.5. Phenotype of the Cluster VI Mutant (ecm33*∆*)*

wall assembly.

(Wald test, FDR = 0.05; Table S6) and were characterized by round mother cells (C115\_A1B and C115\_C), an altered neck position (C105\_A1B and C105\_C), and altered bud direction (C106\_A1B and C106\_C) (Figure S8). In addition, a reduced region of actin at the neck during the M phase (A9\_C) and a lower proportion of cells exhibiting an isotropic pattern of actin (A117) were observed, suggesting that the defects in this mutant manifest before isotropic bud growth (Figure S8A). Consistent with these observations, the bud/mother cell size ratio (C118\_C) (Figure S9) and ratio of cells with a large bud within budded cells (C125\_C; Table S6) were both significantly decreased in *ecm33*∆. We observed a uniform distribution of chitin on the *ecm33*∆ cell surface (Figure 2B). The phosphorylation of Slt2 was increased in *ecm33*∆, suggesting cell wall damage in the cluster VI mutant (Figure 3 The *ecm33*∆ cells exhibited significant differences in 22 morphological parameters (Wald test, FDR = 0.05; Table S6) and were characterized by round mother cells (C115\_A1B and C115\_C), an altered neck position (C105\_A1B and C105\_C), and altered bud direction (C106\_A1B and C106\_C) (Figure S8). In addition, a reduced region of actin at the neck during the M phase (A9\_C) and a lower proportion of cells exhibiting an isotropic pattern of actin (A117) were observed, suggesting that the defects in this mutant manifest before isotropic bud growth (Figure S8A). Consistent with these observations, the bud/mother cell size ratio (C118\_C) (Figure S9) and ratio of cells with a large bud within budded cells (C125\_C; Table S6) were both significantly decreased in *ecm33*∆. We observed a

and Figure S4). These findings suggest possible roles of *ECM33* in bud growth and cell

uniform distribution of chitin on the *ecm33*∆ cell surface (Figure 2B). The phosphorylation of Slt2 was increased in *ecm33*∆, suggesting cell wall damage in the cluster VI mutant (Figures 3 and S4). These findings suggest possible roles of *ECM33* in bud growth and cell wall assembly.

#### *3.6. Phenotype of the Cluster VII Mutant (sag1*∆*)*

The *sag1*∆ cells exhibited significant differences in 28 morphological parameters (Wald test, FDR = 0.05). The sag1∆ mutation caused a smaller cell size at the G1 phase (C11-1\_A, related to C103\_A, C104\_A, and C12-1\_A) (Figure S10A and Table S6). Accordingly, the nucleus was also smaller at the G1 phase in *sag1*∆ cells (D102\_A, D14-1\_A, and D179\_A) (Table S6). We observed the same trend (smaller bud size) at the M phase (C11-2\_C, related to C107\_C, C108\_C, C12-2\_C, C102\_C, and C101\_C) (Figure S10B and Table S6). Moreover, delocalized actin patches were observed frequently in *sag1*∆ cells (A111 and A112) (Figures S10C and S11), suggesting the perturbation of actin polarization and polarized bud growth. The size of the actin region in *sag1*∆ was more heterogeneous at the S/G2 phase (ACV7- 1\_A1B). We observed increased phosphorylation of Slt2 in *sag1*∆, suggesting cell wall damage in the cluster VII mutant (Figure 3 and Figure S4). Although *SAG1* is thought to play an important role only in the mating aggregation process [43,44], this is the first study revealing its effects on cell morphology during the vegetative growth phase.

#### *3.7. Mannoprotein Gene Duplication*

Many mannoprotein genes have been generated by gene duplication (Table S1 and Figure S12). Therefore, the effects of gene duplication were examined by measuring its impact on the morphological phenotype of each mutant (Table S8). More than 80% of the mutants with duplicated genes belonged to clusters III and IV and exhibited no obvious changes in their morphological phenotypes. The remaining *ccw12*∆ (I), *ecm33*∆ (VI), and *tir3*∆ (II) mutants exhibited changes in the morphological phenotype, but no obvious changes were observed in the deletion mutations of their counterparts. This is probably because gene duplication can result in functional bias. On the other hand, among strains with deleted genes unrelated to gene duplication, a significantly lower percentage of the mutants exhibited no obvious changes in the morphological phenotype (Table S8). Taken together, the duplication of mannoprotein genes resulted in a reduction in their functional effects, which made it difficult to examine the morphological phenotype of these gene-deletion strains.

#### *3.8. Comparisons of Morphology and Fitness among Mannoprotein Mutants*

Associations between the comprehensive morphological phenotypes of the 32 mannoprotein mutants and the fitness of these mutants were assessed. Our morphological analysis including 490 morphological parameters revealed 12 mannoprotein mutants with significant abnormalities in at least one morphological parameter (Wald test, FDR = 0.05; Table S6). On the other hand, the fitness analysis of the gene-deleted strains revealed only one mutant (*ccw12*∆) with a significantly decreased growth rate in normal medium (Wald test, FDR = 0.05) (Figure 5A). The *ccw12*∆ mutant exhibited the greatest changes in its morphological phenotype. More differences were found in the morphological phenotype among the mutants than in fitness aspects, probably because of the high sensitivity of morphological phenotyping [12]. The morphological phenotype was also considered to be more greatly affected by the disruption of cell wall proteins.

**Figure 5.** Mannoprotein analyses based on omics studies. (**A**). Scatter plot representing fitness-related defects. The dashed red line indicates a false discovery rate (FDR) of 5%. Data are from Warringer et al. [10]. Fitness data for *ccw14*∆ were not available in the dataset. (**B**). Bar plot showing the average cell wall mannoprotein abundances. Inset: A subset of the data. Data were obtained from [36]. (**C**). Protein–protein interactions among mannoprotein proteins are shown. Black line shows physical interaction. Data are from Oughtred et al. [37]. (**D**). Genetic interactions (GIs) among mannoprotein genes presented as blue (negative GI) or yellow (positive GI) lines (*p* < 0.05). Data are from Costanzo et al. [39]. *EGT2* did not have any significant GIs. In all sections, mutants are color-coded according to Gaussian mixture model clustering of morphological data (see Figure 1).

#### *3.9. Comparisons of Mannoprotein Mutants and Glycosylation-Defective Cells in Morphology*

The remarkable differences in morphological phenotype found for *ccw12*∆ can be explained in terms of protein expression levels (Figure 5B). Yeast cells contain approximately 190,000 Ccw12 protein molecules per cell, accounting for more than 40% of all mannoproteins. The second most highly expressed mannoprotein is Cwp2, with approximately 93,000 molecules expressed per cell. As no morphological abnormalities were detected in *cwp2*∆, the expression level of a mannoprotein originally expressed at a high level would more markedly affect the morphology.

In order to know which cell wall metabolic pathways are relevant to mannoprotein function, we compare the morphology of *ccw12*∆ with those of the cells treated with cell wall agents. For this purpose, we used tunicamycin, echinocandin B, nikkomycin Z, and hydroxyurea, which affect protein glycosylation, 1,3-β-glucan synthesis, chitin synthesis, and DNA replication, respectively (Figure S13). We found that *ccw12*∆ is the most similar to the tunicamycin-treated cells (*r* = 0.813), implying a close relationship between protein glycosylation and mannoprotein function. *ccw12*∆ was also similar to the echinocandin B-treated cells (*r* = 0.723), but not similar to the cells treated with nikkomycin Z (*r* = 0.347) or hydroxyurea (*r* = 0.315). Taken together, these observations indicate that defects in *CCW12* resulted in serious damage to yeast cells, similar to defects in protein glycosylation and 1,3-β-glucan, which is the main filamentous component of the yeast cell wall.

## *3.10. Comparison of Morphological Clustering Results with Those from Analyses of Other Omics Data*

We compared our clustering data with other omics data on interactions. A survey of comprehensive data on PPIs identified only one unidirectional interaction between Pir3 and Cis3 (Figure 5C). However, neither *pir3*∆ nor *cis3*∆ exhibited detectable changes in the morphological phenotype in the present study. Therefore, we could not infer the biological significance of the interaction between these two proteins based on morphological phenotyping. In addition, studying the PPI profile at the proteomic level did not reveal any similar patterns of PPI frequency among members of the same cluster (Figure S14A,B and Table S9). There were no associations between interactome profiles in each cluster either (Figure S14C).

With regard to GIs, we identified 26 positive and 42 negative relationships among the 32 mannoprotein genes. As with the PPI network, GIs among the 32 mannoprotein genes could not be directly linked to molecular functionality (Figure 5D). However, the lack of detectable morphological and fitness defects in many of the individual mannoprotein mutants may be explained by negative GIs comprising more than half (~61.7%) of all GIs. The lack of defects may be due to the existence of parallel pathways with the same or similar biological functions, such as the preservation of the cell wall structure. There were no noticeable GI patterns based on frequency of an interactome profile (Figure S15A,B and Table S10) or correlations between the members of a given cluster (Figure S15C).

Perturbations upon exposure to 106 different chemical compounds were tested in mannoprotein mutants, and the data are summarized in the SGD (Table S11). The chemicalgenetic profiles of the mannoproteins were then visualized as a scatter plot in twodimensional space representing the deletion mutants and chemicals (Figure S16). The comparison of the frequency of each mutant revealed that the chemical response phenotypes of *ccw12*∆ and *ecm33*∆ have been frequently tested. Of the seven clusters, only members of cluster V (*dse2*∆, *egt2*∆, and *sun4*∆) exhibited similar fitness defects with (S)-lactic acid (5.1% *w*/*v*) and miconazole (1000 µg/mL). Otherwise, the results of chemicalgenetic profiling did not appear to be linked to molecular functionality.

Morphological phenotyping of the mannoprotein mutants clearly accentuated unique aspects of the functional network that cannot be identified using other omics technologies. Thus, morphological phenotyping, as a complementary tool, provides deeper knowledge on cell wall organization, remodeling, and protein function. We succeeded in clustering 32 mannoproteins into seven groups based on their morphology and elucidated their specific functions in the cell (Figure 6).

their specific functions in the cell (Figure 6).

**Figure 6**. Schematic representation of the mannoprotein responsibility matrix. Stacked bars show the numbers of disturbed CalMorph parameters (Wald test, FDR = 0.05) related to actin, the cell wall, and the nucleus (illustrated in red, green, and blue, respectively) in mutants of clusters I, II, V, VI, and VII. Prominent implications of morphological defects caused by each mutation are illustrated in a small budding yeast cell where actin, the cell wall, and the nucleus are shown in red, green, and blue, respectively. Green dashed circle represents larger mother cell. Mutants are color-coded according to Gaussian mixture model clustering of morphological data (see Figure 1). **Figure 6.** Schematic representation of the mannoprotein responsibility matrix. Stacked bars show the numbers of disturbed CalMorph parameters (Wald test, FDR = 0.05) related to actin, the cell wall, and the nucleus (illustrated in red, green, and blue, respectively) in mutants of clusters I, II, V, VI, and VII. Prominent implications of morphological defects caused by each mutation are illustrated in a small budding yeast cell where actin, the cell wall, and the nucleus are shown in red, green, and blue, respectively. Green dashed circle represents larger mother cell. Mutants are color-coded according to Gaussian mixture model clustering of morphological data (see Figure 1).

#### **4. Discussion 4. Discussion**

In this study, we used high-dimensional morphological phenotyping to gain a system-level understanding of 32 cell wall mannoproteins in *S. cerevisiae*. We found 12 mannoprotein mutants with significant abnormalities in at least one morphological parameter. Nearly 30% of the 490 unimodal morphological parameters examined were affected in the mannoprotein mutants, implying distinct roles of mannoproteins in cell morphology. Multivariate analysis revealed seven groups of mutants categorized according to the effects of the mutation on their functions. The results indicate that high-dimensional morphological phenotyping of mannoprotein mutants is an effective approach for determining the responsibility matrix of yeast mannoproteins, which is difficult to obtain with other omics technologies. In this study, we used high-dimensional morphological phenotyping to gain a systemlevel understanding of 32 cell wall mannoproteins in *S. cerevisiae*. We found 12 mannoprotein mutants with significant abnormalities in at least one morphological parameter. Nearly 30% of the 490 unimodal morphological parameters examined were affected in the mannoprotein mutants, implying distinct roles of mannoproteins in cell morphology. Multivariate analysis revealed seven groups of mutants categorized according to the effects of the mutation on their functions. The results indicate that high-dimensional morphological phenotyping of mannoprotein mutants is an effective approach for determining the responsibility matrix of yeast mannoproteins, which is difficult to obtain with other omics technologies.

clustering 32 mannoproteins into seven groups based on their morphology and elucidated

#### *4.1. Ccw12 Is a Major Cell Wall Stabilizer 4.1. Ccw12 Is a Major Cell Wall Stabilizer*

The highly pleiotropic morphological defects of *ccw12*Δ cells, including the wide neck, a typical phenotype of cell wall mutants [45], and altered cell shape for both mother and daughter compartments [46], clearly indicated the important role of Ccw12 as a major structural component of the cell wall [47]. This small (133 amino acid residues) and highly glycosylated GPI-anchored protein has been previously shown to impact the maintenance of newly synthesized areas of the cell wall [13] and cell fitness [10]. In addition, *ccw12*∆ has been reported to affect 473 genes acting in various cellular pathways, including 32 genes directly involved in the construction and remodeling of the cell wall [47]. Here, we The highly pleiotropic morphological defects of *ccw12*∆ cells, including the wide neck, a typical phenotype of cell wall mutants [45], and altered cell shape for both mother and daughter compartments [46], clearly indicated the important role of Ccw12 as a major structural component of the cell wall [47]. This small (133 amino acid residues) and highly glycosylated GPI-anchored protein has been previously shown to impact the maintenance of newly synthesized areas of the cell wall [13] and cell fitness [10]. In addition, *ccw12*∆ has been reported to affect 473 genes acting in various cellular pathways, including 32 genes directly involved in the construction and remodeling of the cell wall [47]. Here, we confirmed that *ccw12*∆ cells exhibited the most significant morphological defects with differences found for 81 parameters. Ccw12 is localized at the presumptive budding site, around the bud, and at the septum [47], which explains the defect in the neck width of

*ccw12*∆. An abnormal round cell morphology was also reported previously for *ccw12*∆ cells [13]. The defect in *CCW12* impacted another component of the cell wall because staining using wheat germ agglutinin (WGA) revealed the abnormal localization of chitin. Whereas chitin is located at the budding site in the WT strain, a uniform distribution of chitin on the cell surface was observed in *ccw12*∆ cells. Taken together, these results further confirmed that Ccw12 plays a major role in the maintenance of a rigid cell shape and the stabilization of the cell wall structure.

#### *4.2. Cluster V Member Genes Encode Endoglucanases*

After cytokinesis, mother and daughter cells undergo cell separation, which requires enzymatic digestion of the cell wall [48,49]. Dse2 is a well-known hydrolytic enzyme (glucanase) that functions exclusively in efficient cell separation from the daughter cell side [49,50]. Other cluster V member genes (*EGT2* and *SUN4*) have also been reported to encode glycosidases, and our results clearly showed that genetic perturbation prevented efficient daughter cell separation in all cluster V mutants. Consistent with the mutant phenotypes, co-localization studies have revealed that Dse2, Egt2, and Sun4 form a complex at the birth scar [51]. Cluster V mutants exhibited no defects in cell-cycle progression or daughter cell growth in the next cell cycle, indicating lesser effects of these genes in cell proliferation. Due to redundancy arising from intertwining pathways and the proteins involved, it was not clear how precisely diverse cell wall digestion systems are integrated to achieve effective cell separation; for example, *SUN4* genetically interacts negatively with some septin construction genes, including *CDC11* and *CDC12*, making its role in cell separation complex. Interestingly, our kinetic analysis revealed the temporal order among glucanases/chitinases. Dse2, Egt2, and Cts1 were deposited at the division site after septum synthesis was completed. However, Egt2 preceded Dse2 and Cts1. Therefore, Egt2 may be involved in cell wall maturation and making the wall architecture conducive for cell separation, whereas Dse2 and Cts1 are septum-hydrolyzing enzymes that arrive at the division site during the last step of cell separation. This finding suggests that glucanaseand chitinase-mediated cell separation is accomplished in a stepwise process. Consistent with the above observations, the expression of cell-separation genes is also regulated in a strict temporal order [52], as observed in our kinetic analysis. Early enzymes, such as Egt2, may function to remodel the cell wall or septum structure to facilitate the delivery of Cts1 to the PS [49].

#### *4.3. ECM33 Plays a Role in Bud Growth*

The molecular function of Ecm33 has not been fully elucidated. Previous studies suggested that it may play roles in determining cell shape [7], cell wall biogenesis [53,54], and apical growth [55]. Consistent with those previous reports, we confirmed that the roundness of mother cells (C115) and bud site selection (C106) were perturbed at both the S/G2 and M phases in *ecm33*∆. *ECM33* also has strong negative GIs with mannosyltransferase genes including *MNN11*, *ANP1*, and *HOC1*, which can explain the role of *ECM33* in cell wall assembly.

The smaller proportion of *ecm33*∆ cells exhibiting an isotropic pattern of actin suggests that *ECM33* functions before isotropic bud growth. However, apical bud growth seemed normal because the long and short axis lengths of the buds as well as their ratio were not significantly altered in *ecm33*∆ cells. Therefore, one possibility is that the apical and isotropic bud growth switch is delayed in the mutant. It has also been reported that *ECM33* deletion triggers the activation of the CWI pathway through the phosphorylation of Slt2 [54]. Although the CWI pathway is involved in cell-cycle checkpoints, such as the cell wall integrity checkpoint and cell morphological checkpoint, it is unlikely that any cell-cycle checkpoints were activated because cell-cycle progression appeared to be normal in *ecm33*∆ cells. However, further studies are needed to determine how Ecm33 impacts cell-cycle progression.

#### *4.4. SAG1 Deletion Perturbs Actin Distribution during Vegetative Growth*

*SAG1* (*AGα1*) encodes a cell-adhesion molecule called α-agglutinin in *MATα* cells [42,55], but the function of this molecule during the vegetative growth of *MATa* cells has yet to be identified. In this study, we examined the morphological phenotype of *sag1*∆ *MATa* cells. The results showed that the *sag1*∆ mutation affects the mother and bud cell sizes at the G1 and M stages of the cell cycle, respectively, in *MATa* cells. It also perturbed actin polarization and polarized bud growth. As Sag1 binds directly to Aga1, it would be interesting to investigate the phenotype of *aga1*∆. However, it was difficult to analyze *aga1*∆ because the mutant cells were not suitable for morphological phenotyping due to their propensity to aggregate. The construction of weak alleles of *AGA1* will be necessary to examine the morphological phenotype and investigate its relationship with Sag1.

#### **5. Conclusions**

This study provided a comprehensive analysis of morphological phenotypes of yeast mannoprotein mutants. The morphology of each cluster of mutants could be explained by the molecular functions of the mannoproteins. The cluster I gene (*CCW12*) encodes a mannoprotein that accounts for 40% of the total mannoproteins in a cell, plays a major structural role, and contributes the most to cell morphogenesis. The cluster II genes (*CCW14*, *FLO11*, *SRL1*, and *TIR3*) do not play structural roles but have similar effects on cell size and cell shape. The cluster V genes (*DSE2*, *EGT2*, and *SUN4*) encode glucosidases, which are required for cell separation. The cluster VI gene (*ECM33*) is required for bud growth and cell wall assembly. Finally, the cluster VII gene (*SAG1*) is required for cell aggregation and is important for determining cell size and actin organization. Cluster III and cluster IV genes do not play major roles in cell morphogenesis. The results presented here increase our understanding of the mechanistic and functional roles of glycoproteins in cell morphogenesis. Morphology-based analysis seems to be a practical means of relating morphological defects to underlying molecular mechanisms, indicating the sensitivity of our approach for determining the responsibility matrix of mannoproteins regarding maintaining the cell wall structure.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/jof7090769/s1, Figure S1. Multivariate analysis of mannoprotein morphological data, Figure S2. Specific morphological features of ccw12∆ cells (cluster I), Figure S3. Specific morphological features of ccw12∆ buds (cluster I), Figure S4. Complete gel of western blotting of phosphorylated Slt2, Figure S5. Specific morphological features of mother cells of all mutants in cluster II, Figure S6. Specific morphological features shared among members of cluster V, Figure S7. Morphological defects in cluster V do not affect the cell cycle, Figure S8. Specific morphological features of ecm33∆ (cluster VI), Figure S9. Morphological parameters related to cell size in ecm33∆ (cluster VI) versus other mannoprotein mutants, Figure S10. Specific morphological features of sag1∆ (cluster VII), Figure S11. Specific actin-related morphological features of sag1∆ (cluster VII) in other mutants, Figure S12. Gene homology among mannoproteins, Figure S13. Morphological similarity of mannoproteins and drugtreated wild-type cells, Figure S14. Protein–protein interaction (PPIs) network, Figure S15. Genetic interaction network, Figure S16. Chemical-genetic profile of mannoproteins, Table S1. List of 36 cell wall mannoprotein strains, Table S2. List of strains to study kinetics of mannoproteins (Dse2 and Egt2; cluster V) and proteins involved in the cytokinesis and cell separation, Table S3. Oligonucleotides (A) and plasmids (B) used in this study, Table S4. Parameters showed statistically significant difference at least in one mutant compared with the null distribution (Wald test, FDR = 0.05), Table S5. Significant loadings of first five PC spaces (used for GMM clustering) after Bonferroni correction (*t*-test, *p* < 0.05), Table S6. Significant parameters of each mannoprotein mutant (Wald test, FDR = 0.05), Table S7. Pearson correlation coefficient (r)1 between Dse2-GFP and other proteins, Table S8. Duplicated and non-duplicated genes of mannoproteins considering their functional responsibility and diversification, Table S9. Protein-Protein interactions of 32 mannoproteins. Data obtained from Oughtred et al. (2019), Table S10. Genetic interactions of 32 mannoproteins. Data obtained from Costanzo et al. (2016), Table S11. Chemical-genetic profile of 32 mannoprotein mutants. Data obtained from the *Saccharomyces* Genome Database (SGD).

**Author Contributions:** Conceptualization, F.G. and Y.O.; Experiments, Y.L., K.I.-N., H.O. and S.Y.; Methodology, F.G. and S.O.; Formal Analysis, F.G. and S.O.; Data Curation, A.S., E.B. and S.Y.; Writing—Draft Preparation, F.G., H.O. and Y.O.; Writing—Review and Editing, Y.O., F.G., E.B. and S.Y.; Visualization, F.G.; Supervision, E.B. and Y.O.; Project Administration, Y.O. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was supported by Grants-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology, Japan to Y.O. (19H03205) and a MEXT scholarship to F.G. (160693) as well as a National Institutes of Health grant to E.B. (GM115420).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Any additional data will be available upon request to the corresponding author.

**Acknowledgments:** We thank Kuninori Suzuki and other members of the Laboratory of Signal Transduction for their participation in helpful discussions.

**Conflicts of Interest:** The authors declare no conflict of interest.

## **References**


**Marina Valente Navarro <sup>1</sup> , Yasmin Nascimento de Barros <sup>2</sup> , Wilson Dias Segura <sup>2</sup> , Alison Felipe Alencar Chaves <sup>3</sup> , Grasielle Pereira Jannuzzi <sup>4</sup> , Karen Spadari Ferreira <sup>2</sup> , Patrícia Xander <sup>2</sup> and Wagner Luiz Batista 1,2,\***


**Abstract:** Dimorphic fungi of the *Paracoccidioides* genus are the causative agents of paracoccidioidomycosis (PCM), an endemic disease in Latin America with a high incidence in Brazil. This pathogen presents as infective mycelium at 25 ◦C in the soil, reverting to its pathogenic form when inhaled by the mammalian host (37 ◦C). Among these dimorphic fungal species, dimorphism regulating histidine kinase (Drk1) plays an essential role in the morphological transition. These kinases are present in bacteria and fungi but absent in mammalian cells and are important virulence and cellular survival regulators. Hence, the purpose of this study was to investigate the role of PbDrk1 in the cell wall modulation of *P. brasiliensis*. We observed that PbDrk1 participates in fungal resistance to different cell wall-disturbing agents by reducing viability after treatment with iDrk1. To verify the role of *PbDRK1* in cell wall morphogenesis, qPCR results showed that samples previously exposed to iDrk1 presented higher expression levels of several genes related to cell wall modulation. One of them was *FKS1*, a β-glucan synthase that showed a 3.6-fold increase. Furthermore, confocal microscopy analysis and flow cytometry showed higher β-glucan exposure on the cell surface of *P. brasiliensis* after incubation with iDrk1. Accordingly, through phagocytosis assays, a significantly higher phagocytic index was observed in yeasts treated with iDrk1 than the control group, demonstrating the role of PbDrk1 in cell wall modulation, which then becomes a relevant target to be investigated. In parallel, the immune response profile showed increased levels of proinflammatory cytokines. Finally, our data strongly suggest that PbDrk1 modulates cell wall component expression, among which we can identify β-glucan. Understanding this signalling pathway may be of great value for identifying targets of antifungal molecular activity since HKs are not present in mammals.

**Keywords:** histidine kinase; dimorphism; *Paracoccidioides*; paracoccidioidomycosis; cell wall

## **1. Introduction**

Paracoccidioidomycosis (PCM) is a systemic granulomatous human disease endemic in Latin America. It is caused by *Paracoccidioides* spp., a thermally-dimorphic fungus that presents as an infective mycelium in the environment, and it switches to a pathogenic yeast form in the mammalian host [1,2]. Its clinical manifestations occur in two distinct forms, acute or subacute and chronic [3], affecting mainly the lungs, but it is capable of spreading to other tissues [4,5]. Primary infection usually occurs by inhaling propagules (conidia) produced during the mycelial form [6]. Once inhaled, fungal propagules will

**Citation:** Navarro, M.V.; de Barros, Y.N.; Segura, W.D.; Chaves, A.F.A.; Jannuzzi, G.P.; Ferreira, K.S.; Xander, P.; Batista, W.L. The Role of Dimorphism Regulating Histidine Kinase (Drk1) in the Pathogenic Fungus *Paracoccidioides brasiliensis* Cell Wall. *J. Fungi* **2021**, *7*, 1014. https://doi.org/10.3390/jof7121014

Academic Editors: María Molina and Humberto Martín

Received: 30 September 2021 Accepted: 24 November 2021 Published: 26 November 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

be recognized by cells of the innate immune system [7]. The recognition of fungal cell wall components begins with pathogen-associated molecular patterns (PAMPs) through pathogen recognition receptors (PRRs). These receptors include Toll-like receptors (TLRs), mannose receptors, complement pathway molecules and lectin family receptors (CLRs), such as dectin-1 [8]. The interaction of these molecules with fungal yeasts leads to activation of the innate immune response, consequently activating mediators involved in eliminating these pathogens and controlling the adaptive immune response [9].

However, fungi have mechanisms to prevent their elimination by the host's immune system [10]. Several fungi that engage in morphological transitions are of great medical importance, such as *Talaromyces marneffei* (*Penicillium marneffei*), *Blastomyces dermatitidis*, *Coccidioides immitis*, *Histoplasma capsulatum*, *Sporothrix schenckii*, and *Paracoccidioides* spp. [11]. The transition in *Paracoccidioides* spp. and other dimorphic fungi is essential for the establishment of the disease [12]. This alters not only the cell morphology but also the composition of the cell wall elements. In *Paracoccidioides* spp. mycelium, there is a prevalence of β-1,3-glucan and β-1,6-glucan, and in the yeast form, there is a prevalence of α-1,3-glucan and chitin [13]. This ensures fungal survival in the host environment since the content of α-1,3-glucan is correlated with the degree of fungal virulence [14]. In addition, masking the presence of β-1,3-glucan molecules, a highly immunogenic structure, is recognized by the dectin-1 receptor of phagocytic cells [15]. The morphological switch is believed to be an additional evasion strategy against phagocytic cells and mechanisms for recognition of the cell wall components [15]. In this context, the fungal cell wall plays an important role in immunological recognition.

In *Paracoccidioides* spp., different genes are expressed according to the phase (yeast or mycelium) [16]. The mechanism of this transition has been unclear. However, genes related to the control of the mycelium-yeast transition (M-Y) have recently been identified in *B. dermatitidis* and *H. capsulatum*, including dimorphism-regulating histidine kinase (*DRK1*). *DRK1* is mainly expressed in the yeast phase [11] of *B. dermatitidis* [17], *S. schenckii* [18], and *T. marneffei* [19] and, more recently, it was characterized in *P. brasiliensis* [20].

Histidine kinases (HKs) were discovered in the 1980s in *Escherichia coli* [21], and were believed to be present only in bacteria. In the 1990s, they were also discovered in plants, fungi, archaea, cyanobacteria and amoebas [22]. In fungi, the functions attributed to HKs have not been explored very well [23]. HKs are classified based on phylogenetic analyses; in fungi, there are 16 groups, determined by the C- and N-terminal regions and the domains present in each group [22,24,25]. This is a signal transduction mechanism that contains a conserved kinase domain and a conserved regulatory domain. After an extracellular stimulus, the HK domain is autophosphorylated on a histidine residue, followed by a phosphate group transfer to the regulatory domain in an aspartate residue, which catalyses a downstream reaction of the effector domain that leads to downstream signalling [26].

Among the characterized pathways in fungi, we can mention the response to osmotic stress [25], oxidative protection against phagocytic cells [27] and regulation of the dimorphism in pathogenic fungi [17,18,20,28]. In *P. brasiliensis*, *DRK1* is known to be a group III histidine kinase essential to the dimorphic transition process [20].

Since HKs are known to regulate morphological switches in *P. brasiliensis*, this work aimed to characterize the PbDrk1 protein, which is involved in the transition from mycelium to yeast. This investigation is of great interest since HKs use a phosphorylation mechanism where the amino acid phosphoryl-receiving groups are aspartate and histidine residues, unlike the serine, threonine and tyrosine residues that are prevalent in mammals. Thus, knowing that these molecular sensors are absent in humans, it is extremely important to study the components that are part of these activation pathways, as they may represent potential molecular targets in the development of new antifungal agents.

#### **2. Materials and Methods**

#### *2.1. Fungal Isolates and Growth Conditions*

The *P. brasiliensis* isolate Pb18 was grown in yeast peptone dextrose modified medium (mYPD) (0.5% yeast extract, 1% peptone, and 0.5% glucose, pH 6.7) for 4 to 5 days at 37 ◦C and shaking at 150 rpm. For mycelium growth, viable yeast cells were cultivated in mYPD at 25 ◦C for 7 days at 150 rpm. Viability was assessed by Trypan blue 0.4% counting on Neubauer's chamber, using the formula: cell viability(%) = viable cells /total cells × 100. All chemicals were purchased from Sigma-Aldrich (St. Louis, MO, USA) unless otherwise mentioned.

#### *2.2. Histidine Kinase Inhibitor Susceptibility*

About 1 <sup>×</sup> <sup>10</sup><sup>6</sup> yeast were incubated with different concentrations (100, 50, 25, 12.5, and 6.25 µg/mL) of Fludioxonil (Thermo Scientific, Waltham, MA, USA), a specific inhibitor of class III histidine kinase (iDrk1). The inhibitor was solubilized in DMSO (dimethylsulfoxide). Yeasts were incubated for 24 h under constant agitation of 180 rpm at 37 ◦C. Each yeast culture was diluted (10, 50, 100, 500, and 1000 times) in YPDmod broth, and 10 µL of each suspension was plated to YPDmod agar medium. Plates were photographed after 7 days of growth at 37 ◦C. This assay was performed in biological triplicate.

#### *2.3. Dimorphic Transition Assay*

Yeast cells of Pb18 were grown in mYPD agar (pH 6.5) at 37 ◦C for 4 to 5 days and inoculated in mYPD broth medium (pH 6.5). Then yeasts were incubated at 25 ◦C for five to six days to reverse yeast to mycelium entirely. After the complete transition, yeasts were centrifuged at 3000× *g* and washed with PBS buffer (pH 7.2). The mycelium was then seeded in 6-well plates and 20 µM iDrk1 (Fludioxonil) was added. The samples were monitored every 24 h under an optical microscope (Zeiss) at 100× magnification. Every 24 h, the culture medium was supplemented with 20 µM fludioxonil (because of inhibitor photodegradation). With each addition, a new solution was prepared to guarantee its activity. This assay was performed in a biological duplicate.

#### *2.4. Cell Wall Disturbing Agents Spot Test*

The sensitivity of Pb18 to cell wall disruptors was investigated using the spot assay. About 1 <sup>×</sup> <sup>10</sup><sup>6</sup> yeasts were incubated with iDrk1 (25 <sup>µ</sup>g/mL) for 24 h at 37 ◦C at 150 rpm. Each yeast culture was diluted (10, 50, 100, and 500 times) in YPDmod broth, and 10 µL of each suspension was applied to mYPD agar medium supplemented with different cell wall disrupting agents, such as: Congo Red (Congo Red) (2.5 µM), Calcofluor White (1 µg/mL) and sodium chloride (150 mM). The plates were incubated for seven days at 37 ◦C, and then photographed. This assay was performed in biological triplicate.

#### *2.5. RNA Extraction and Real-Time Quantitative PCR Analysis*

Pb18 yeasts were grown for four to five days in YPDmod pH 6.5 medium at 37 ◦C and 150 rpm, counted and the volume was adjusted to 30 mL at a concentration of <sup>1</sup> <sup>×</sup> <sup>10</sup><sup>6</sup> cells/mL. Then, yeasts were incubated with 25 µg/mL of iDrk1 for 24 h at 37 ◦C. After this incubation period, RNA extraction was performed. Samples were centrifuged at 3000× *g* for 10 min at 4 ◦C and washed 3 times with PBS (pH 7.2). Then, in 15 mL tubes, approximately 500 µL of glass beads (425–600 µm—Sigma-Aldrich, San Louis, MO, USA) and 1.5 mL of TRizol® (Invitrogen, Waltham, MA, USA) were added to the sample. The tubes were vigorously vortexed for 6 cycles of 1 min, alternating with 1 min on ice. RNA extraction was performed as previously described [29]. Quantification was performed using spectrophotometry (NanoDrop 2000/2000c, Thermo Fisher Scientific, Waltham, MA, USA). For complementary DNA (cDNA) synthesis, 500 ng of RNA was initially submitted to the DNase I enzyme (Thermo Fisher Scientific, Waltham, MA, USA) and then to ProtoScript First Strand cDNA Synthesis kit (New England BioLabs, Ipswich, MA, USA), according to the manufacturer's instructions. To assess gene expression by

real-time quantitative PCR, the reaction was performed with SYBR® Green Master Mix (Applied Biosystems, Foster City, CA, USA) according to the manufacturer's instructions. The endogenous expression genes for ribosomal protein 60S L34 (L34r) and 18S were used as normalizing controls. For each gene of interest and normalizer gene, a negative reaction control was also added. The samples were prepared in triplicate to a 96-well plate (0.2 mL MicroAmp™ Optical 96-Well Reaction Plate—Applied Biosystems) compatible with the equipment used, and the plate was sealed with an optical adhesive (MicroAmp™ Optical Adhesive Film—Thermo Fisher Scientific, Waltham, MA, USA). The equipment used was the ABI StepOne Plus Real-Time PCR System (Applied Biosystems) with the following conditions: 10 min at 95 ◦C, followed by 40 cycles of 15 s at 95 ◦C and 1 min at 60 ◦C. The dissociation curve included an additional cycle of 15 s at 95 ◦C, 20 s at 60 ◦C, and 15 s at 95 ◦C. The curves of oligonucleotides efficiency were evaluated from a cDNA obtained previously and serially diluted (100, 10, 1, and 0.1 ng/µL). The Ct values of each dilution point were determined and used to make the standard curve and finally calculate the primer efficiency (E = 10(−1⁄slope)−<sup>1</sup> <sup>×</sup> 100). The relative expression was determined based on the 2−∆∆Ct method [30]. The sequences used for each gene are listed in Table 1.

**Table 1.** Oligonucleotides used for real-time quantitative PCR analysis.


#### *2.6. Quantification of Cell Wall Components*

About 1 <sup>×</sup> <sup>10</sup><sup>6</sup> yeast cells were grown and incubated for 24 h with 25 <sup>µ</sup>g/mL of iDrk1 in mYPD at 37 ◦C under stirring at 180 rpm. Then, yeasts were collected, homogenized in blocking solution (0.5% BSA, 5% rabbit serum, 5 mM EDTA, 2 mM NaN<sup>3</sup> in PBS, pH 7.2) and incubated for 30 min at room temperature. Then, yeasts were incubated with 1 µg/mL of the β-glucan binding probe for 1 h on ice. This probe corresponds to the human Dectin-1 receptor fused to the FC portion of mouse IgG1 (Sino Biological, Beijing, China). Samples were incubated with Alexa-488-conjugated anti-mouse IgG secondary antibody (Molecular Probes, Eugene, OR, USA) at a 1:200 ratio for 45 min on ice. Between each incubation, step cells were washed three times with wash buffer (0.5% BSA, 5 mM EDTA, 2 mM NaN<sup>3</sup> in PBS, pH 7.2). Determination of chitin oligomers was performed with the WGA (wheat germ agglutinin) marker conjugated to FITC (Sigma-Aldrich, San Louis, MO,

USA), at a concentration of 25 µg/mL in 1 mL of PBS (pH 7.2). Mannan determination was performed with FITC-conjugated Concanavalin A (Sigma-Aldrich, San Louis, MO, USA), at a concentration of 25 µg/mL in 500 µL of PBS (pH 7.2). Both samples were incubated for 1 h protected from light under agitation at 800 rpm. Next, samples were centrifuged at 3000× *g* for 5 min at 4 ◦C and washed three times with 1 mL of PBS (pH 7.2). Finally, all samples were homogenized in 500 µL of PBS (pH 7.2) and analyzed by flow cytometry (BD FACSCaliburTM, Becton Dickinson) using the FL-1 detection channel. Each sample was prepared in experimental triplicate. A total of 10,000 events were counted and quantification graphs were generated from the median fluorescence intensity (MFI). The data obtained were analyzed using FlowJo software version 10.6.2 (FlowJo, LLC, FlowJo™, Ashland, OR, USA) [31].

#### *2.7. Confocal Microscopy*

To evaluate the exposion of β-glucan molecules through fluorescence confocal microscopy, cells were prepared as described on the previous section and then centrifuged and homogenized in PBS:glycerol (3:1). The slides were prepared with 30 µL of the sample, sealed and analyzed under a confocal microscope (SP8 Lightning, Leica Microsystems). Fluorescence intensity was quantified using ImageJ analysis software (version 1.53i) through corrected total cell fluorescence (CTCF).

#### *2.8. PKA Activity and cAMP Quantification*

PKA activity was performed using the PKA Colorimetric Activity Kit (Thermo Fisher Scientific, Waltham, MA, USA), according to the manufacturer's specifications. For this purpose, 1 <sup>×</sup> <sup>10</sup><sup>7</sup> Pb18 cells were incubated with or without 25 <sup>µ</sup>g/mL of iDrk1 or 1 mM H2O<sup>2</sup> for 30 min in YPDmod at 37 ◦C and 180 rpm. Yeasts were collected and washed with PBS (pH 7.2), centrifuged for 3000× *g* for 10 min and homogenized in Tris-based lysis buffer contained in the kit. Lysis was assessed by adding glass beads and vigorous vortexing (5 cycles of 1 min interspersed with 1 min of incubation on ice). Samples were then centrifuged and the supernatant was recovered. For PKA activity assay, 1 mg of protein was used. Intracellular levels of cyclic AMP (cAMP) were quantified by the Cyclic AMP ELISA Kit (Cayman Chemical, Ann Harbor, MI, USA). Pb18 yeasts were prepared as described above. Cellular lysis was performed according to the manufacturer's specifications, using 10 mM HCl and glass beads. For cAMP quantification, it was used about 0.5 µg of protein. The total protein extracts quantification was performed by Bradford assay [32]. Absorbance was read at 405 nm (BioTek—Synergy HT) and data were plotted in triplicate from the standard curves absorbance values.

#### *2.9. Glycogen Accumulation*

A 10 mL suspension of 5 <sup>×</sup> <sup>10</sup><sup>6</sup> Pb18 yeasts/mL was incubated with or without 25 µg/mL of iDrk1 for 24 or 48 h, followed by centrifugation at 2500 rpm for 3 min (the supernatant was discarded). The pellet was homogenized in 1 mL of iodine solution (0.2% iodine and 0.4% potassium iodide) and incubated for 3 min at room temperature. The samples were again centrifuged at 2500 rpm for 3 min, the supernatant was discarded and the pellet homogenized in 30 µL of the iodine solution. The samples were plated to a 96-well plate, 200 µL of PBS (pH 7.2) was added to each well and the samples were photographed [33].

#### *2.10. Phagocytosis Assay*

In vitro phagocytosis was performed with the J774 macrophage cell line. About 2.5 <sup>×</sup> <sup>10</sup><sup>5</sup> viable cells were plated in a 24-well plate containing RPMI (Gibco, Gaithersburg, MD, USA) supplemented with 10% FBS. A 15 mm diameter circular sterilized coverslip was added to each well. After adhesion, macrophages were primed with 100 ng/mL of LPS 30 min before interaction with *P. brasiliensis*. J774 cell line is activated by LPS showing changes in morphology, such as cytoplasm expansion, contributing to a better

performance in phagocytosis assays [34,35]. The interaction was carried out in a 2:1 ratio (yeast:macrophages) and incubated for 24 h at 37 ◦C and 5% CO2. Previously, yeasts were incubated in the presence or absence of iDrk1 25 µg/mL for 24 h at 37 ◦C. After the interaction, each well was washed with sterile PBS (pH 7.2), coverslips were stained with hematology dyes (Newprov, Paraná, Brazil) and the supernatant was recovered for cytokine assay. The phagocytic index was determined from the protocol established by Popi et al. [36]. After 48 h, wells intended to quantify colony-forming units (CFU) were washed with 1 mL of ice-cold sterile ultrapure water and vigorously homogenized. The supernatant was plated in BHI medium supplemented with 10% FBS, and incubated for seven days at 37 ◦C. After growth, colony-forming units were counted. This experiment was carried out in biological triplicate.

The phagocytosis assay was also assessed by flow cytometry. Samples were prepared as described above. Prior to interaction, yeasts were labeled with CFSE (CellTrace™ CFSE Cell Proliferation—Thermo Fisher). For each 1 <sup>×</sup> <sup>10</sup><sup>7</sup> Pb18 yeasts cells, 3 <sup>µ</sup>L of CFSE (3 µg/µL) were added, with final volume of 500 µL of PBS (pH 7.2) and 0.5% BSA. Yeasts were incubated for 10 min at 37 ◦C, then centrifuged at 3000× *g* for 5 min and washed three times with PBS (pH 7.2). After 24 h, the samples were washed and the cells were detached from the wells using a cell scraper and homogenized in 500 µL of PBS (pH 7.2). Data acquisition was immediately performed in a flow cytometer (BD FACSCaliburTM, Becton Dickinson) using the FL-1 detection channel. After the first acquisition, 8 µL of Trypan Blue 0.4% was added to each sample to quench the signal of yeasts that could be adhered to the cell surface instead of internalized and again, the data was acquired. A total of 10,000 events were counted for each sample and the data were analyzed using the FlowJo software version 10.6.2 (FlowJo, LLC, FlowJo™, Ashland, OR, USA) [37].

#### *2.11. Cytokines Determination*

Supernatants from the phagocytosis assay were collected. The cytokines TNFα and IL12p70 were measured using the DuoSet ELISA kits (R&D Systems, Mineápolis, MN, EUA). The assay was performed in 96-well EIA/RIA plates according to the manufacturer's specifications with modifications. First, plates were coated with 50 µL of capture antibody and incubated at room temperature for 16 h. Wells were blocked with 200 µL of diluent solution (1% BSA in PBS pH 7.2) and incubated at room temperature for 1 h. Then, 50 µL of culture supernatant were added, in triplicate, and 50 µL of the cytokine standard to perform the standard curve. Plates were then incubated for 2 h at room temperature. Next, 50 µL of Streptavidin-HRP solution (provided in the kit) were added to each well and the plate was incubated for 20 min protected from direct light. Finally, Tetramethylbenzidine (TBM) substrate was added and the plates were incubated for 20 min at room temperature, protected from light. At the end of the incubation, 50 µL of stop solution (2N H2SO4) was added. Between each incubation step, the wells were washed three times with 200 µL of wash buffer (0.05% Tween-20 in PBS pH 7.2). Absorbance was read at 430 nm (BioTek— Synergy HT).

#### *2.12. Statistical Analysis*

The data contained in this work were validated with the reproducibility of at least three independent experiments. For comparison analysis, a Student's t-test and significance analysis were performed, as were the one-way variance (ANOVA), followed by Tukey's test. Differences were considered significant when *p* < 0.05.

#### **3. Results**

#### *3.1. Susceptibility of P. brasiliensis to Drk1 Pharmacological Inhibitors*

Studies analysing the role of Drk1 in fungi [17,20] demonstrated that the use of specific inhibitors of group III histidine kinases (iprodione or fludioxonil) are efficient in promoting biological responses. Fludioxonil is a product derived from pyrrolnitrine, a compound isolated from *Pseudomonas pyrrocinia* [38]. The use of this inhibitor is already well established in studies with pathogenic fungi [17,39–42]. Initially, a susceptibility test was assessed with the Pb18 isolate. For this purpose, the yeasts were incubated for 24 h with different concentrations (100 to 6.25 µg/mL) of iDrk1 (fludioxonil) and then inoculated in mYPD medium. After 7 days of incubation, it was observed that there was no reduction in fungal viability at the concentrations tested (Figure 1A). Thus, based on the literature data [17], which demonstrated that a concentration of 25 µg/mL was able to inhibit the activity of Drk1 in *Blastomyces dermatitidis*, this concentration was established for the following experiments (Figure 1A). established in studies with pathogenic fungi [17,39–42]. Initially, a susceptibility test was assessed with the Pb18 isolate. For this purpose, the yeasts were incubated for 24 h with different concentrations (100 to 6.25 µg/mL) of iDrk1 (fludioxonil) and then inoculated in mYPD medium. After 7 days of incubation, it was observed that there was no reduction in fungal viability at the concentrations tested (Figure 1A). Thus, based on the literature data [17], which demonstrated that a concentration of 25 µg/mL was able to inhibit the activity of Drk1 in *Blastomyces dermatitidis*, this concentration was established for the following experiments (Figure 1A).

and the plate was incubated for 20 min protected from direct light. Finally, Tetramethylbenzidine (TBM) substrate was added and the plates were incubated for 20 min at room temperature, protected from light. At the end of the incubation, 50 µL of stop solution (2N H2SO4) was added. Between each incubation step, the wells were washed three times with 200 µL of wash buffer (0.05% Tween-20 in PBS pH 7.2). Absorbance was read

The data contained in this work were validated with the reproducibility of at least three independent experiments. For comparison analysis, a Student's t-test and significance analysis were performed, as were the one-way variance (ANOVA), followed by

Studies analysing the role of Drk1 in fungi [17,20] demonstrated that the use of specific inhibitors of group III histidine kinases (iprodione or fludioxonil) are efficient in promoting biological responses. Fludioxonil is a product derived from pyrrolnitrine, a compound isolated from *Pseudomonas pyrrocinia* [38]. The use of this inhibitor is already well

*J. Fungi* **2021**, *7*, 1014 7 of 21

Tukey's test. Differences were considered significant when *p* < 0.05.

*3.1. Susceptibility of P. brasiliensis to Drk1 Pharmacological Inhibitors* 

at 430 nm (BioTek—Synergy HT).

*2.12. Statistical Analysis* 

**3. Results** 

**Figure 1.** (**A**) Susceptibility of *P. brasiliensis* yeast cells to Drk1 pathway inhibitor (iDrk). A total of 1 × 106 cells/mL were incubated with different concentrations of iDrk1 and incubated for 24 h. Then, the yeasts cells were diluted, plated in solid mYPD medium and incubated in for 7 days at 37 °C. (**B**) Dimorphic transition assay of *P. brasiliensis* in the presence or not of iDrk1. The fungus was cultivated as mycelial, at 25 °C and 150 rpm. Subsequently, it was plated in 6-well plates and incubated at 37 °C in mYPD medium supplemented or not with 20 µM iDrk1. Cultures were monitored every 24 h and observed under an optical microscope to evaluate the dimorphic transition. **Figure 1.** (**A**) Susceptibility of *P. brasiliensis* yeast cells to Drk1 pathway inhibitor (iDrk). A total of 1 <sup>×</sup> <sup>10</sup><sup>6</sup> cells/mL were incubated with different concentrations of iDrk1 and incubated for 24 h. Then, the yeasts cells were diluted, plated in solid mYPD medium and incubated in for 7 days at 37 ◦C. (**B**) Dimorphic transition assay of *P. brasiliensis* in the presence or not of iDrk1. The fungus was cultivated as mycelial, at 25 ◦C and 150 rpm. Subsequently, it was plated in 6-well plates and incubated at 37 ◦C in mYPD medium supplemented or not with 20 µM iDrk1. Cultures were monitored every 24 h and observed under an optical microscope to evaluate the dimorphic transition.

#### *3.2. Role of PbDrk1 in P. brasiliensis Cell Wall Maintenance*

The transition from mycelium to yeast triggers the cell wall morphogenesis machinery, which involves synthesizing several cell wall sugars and proteins critical to survival during infection and evasion of the immune system [43]. In *Penicillium marneffei* pathogenic fungus, the role of *DRK1* is essential for stress adaptation, hyphal morphogenesis, and cell wall integrity [44]. Previously, we showed *PbDRK1* transcription to be phase-specific for the yeast form and demonstrated that PbDrk1 participates in dimorphic switching in *P. brasiliensis* when iprodione (another Drk1 inhibitor) is used [20]. To confirm the action of fludioxonil (iDrk1) in the *P. brasiliensis* yeast-mycelium switch, a dimorphic transition assay was performed. Initially, the fungus was cultivated in the mycelial form at 25 ◦C and then it was incubated at 37 ◦C in the presence or absence of iDrk1. The dimorphic transition was then followed every 24 h under an optical microscope. After 24 h at 37 ◦C, it was possible to observe the formation of yeasts in the distal portion of the hyphae in the control group, and at 96 h, there was a predominance of yeasts. In the group treated with iDrk1, at 96 h, there was a predominance of hyphae. Thus, the addition of fludioxonil also impairs the fungus's ability to make the complete transition from mycelium to yeast (Figure 1B).

In addition to the already known mechanism involving the dimorphic transition [17,18,20,45], it was observed that the strain deleted for Drk1 presents with sensitivity for conidia germination [44]. Based on this evidence, Pb18 yeasts were treated with iDrk1 and inoculated in YPDmod medium containing cell wall stressing agents such as Congo red (CR), Calcofluor White (CFW), and sodium chloride (Figure 2A). CR and CFW dyes are classically used in

studies involving the synthesis and organization of fungal cell wall [46]. Both molecules have two groups of sulfonic acids that exert antifungal activity [47]. The action of CFW and CR occurs through binding to nascent chitin chains, preventing the access of enzymes that promote the binding of chitin with β-1-3-glucan and β-1-6-glucan chains. As a result, the cell wall becomes weakened, which can compromise its viability [47]. Sodium chloride promotes osmotic stress and changes the structure of the cell wall [48]. After seven days of incubation, it was observed that, under all conditions, previous exposure to iDrk1 induced a fairly dramatic viability reduction, especially under osmotic stress (Figure 2A). Previous data from our group [20] showed an increase in the number of *PbDRK1* transcripts when the fungus is subjected to osmotic stress. The data show that the inhibition of Drk1 results in an increased sensitivity to cell wall stressors and to osmotic stress.

#### *3.3. Modulation of Cell Wall Gene Expression in P. brasiliensis*

Previous data demonstrated that *PbDRK1* is mainly expressed in the yeast phase [20]. However, the associated pathways remain poorly understood. Therefore, to investigate the possible targets regulated by *PbDRK1*, Pb18 yeasts were incubated in the absence or presence of iDrk1 for 24 h. Then, the RNA extraction protocol was applied, followed by a cDNA synthesis reaction. Finally, RT-qPCR analysis of several genes involved in cell wall synthesis was performed: *CHT3*, *CHS2*, *CHS3*, *CHS4*, and *CHS5* (genes involved in the synthesis and maintenance of chitins) and *FKS1*, *KRE6*, *PHR2*, *GEL3*, and *AGN1* (genes involved in glucan synthesis and maintenance).

As shown in Figure 2B, genes encoding chitin synthase enzymes *CHT3*, *CHS2* and *CHS3* exhibited a 3.7-, 5.8- and 2.0-fold increase, respectively, in samples treated with iDrk1. However, there were no significant changes in the *CHS4* and *CHS5* genes (Figure 2B). On the other hand, the *CHS4* and *CHS5* genes were more highly expressed in the mycelial phase of Pb18 [49]. Thus, these data suggest that iDrk1 may modulate the expression of some chitin synthesis genes.

Significant increases in the transcript levels of several genes related to β-glucan synthesis and cell wall integrity have been observed (Figure 2C). A 4-fold increase in the *FKS1* gene, which encodes a β-(1,3)-glucan synthase, was observed. The *KRE6* gene showed a 2.7-fold increase in expression levels when compared with the no-treatment control. This gene is important in the β-(1,6)-glucan synthesis process and for molecules that are part of the β-(1,3)-glucan net composition [13,50]. The *PHR2* gene showed a 5.5-fold increase in expression levels (Figure 2C). This gene is involved in the maintenance of the cell wall and fungal virulence [51]. Finally, the expression of the *GEL3* gene exhibited a fourfold increase compared to the control without treatment. This gene is part of the process of elongation of the β-(1,3)-glucan chains and cell wall integrity [52,53]. On the other hand, the *AGN1* gene, which is involved in the maintenance and synthesis of α-(1,3)-glucan and highly expressed in the yeast phase [54], showed no difference in transcript levels (Figure 2C). These findings support the hypothesis that PbDrk1 could regulate negatively genes involved with the cell wall synthesis.

#### *3.4. Modulation of the Cell Wall Components*

As previously demonstrated, the inhibition of PbDrk1 was responsible for modulating the expression of genes involved in the synthesis of important cell wall components, such as *FKS1*. Associated with this, we observed a greater sensitivity of the fungus to cell wall disturbances when treated with iDrk1. Therefore, we decided to evaluate the levels of β-glucan, chitin and mannan in the Pb18 cell wall after treatment with iDrk1. For this purpose, β-glucan labelling was performed using the Dectin-1-Fc probe. Next, chitin levels were determined using the WGA (wheat germ agglutinin) marker conjugated with FITC [55]. This lectin molecule has an excellent affinity for N-acetyl-β-D-glucosaminyl residues and N-acetyl-β-D-glucosamine oligomers. Finally, to verify the mannan levels, concanavalin A conjugated with FITC was used [31]. This molecule has a high affinity for terminal residues of α-D-mannose and α-D-glucose.

**Figure 2.** (**A**) Growth of *P. brasiliensis* in the presence of cell wall disrupting agents. A total of 1x106 cells/mL were subjected to 25 µg/mL of iDrk1 and incubated for 24 h. Next, yeast cells were diluted, plated in solid mYPD medium containing different agents that disturb the cell wall, such as Calcofluor White (CFW), Congo Red and NaCl. Finally, cells were incubated in for seven days at 37 °C. Expression of cell wall morphogenesis related genes in *P. brasiliensis*. Pb18 cells were subjected to 25 µg/mL of iDrk1 for 24 h and then total RNA extraction was performed. The related genes (**B**) synthesis of chitin, such as *CHT3*, *CHS2*, *CHS3*, *CHS4* and *CHS5* and (**C**) synthesis of glucans such as *FKS*, *KRE6*, *PHR2*, *GEL3* and *AGN1* were analyzed, where *p*-value ≤ 0.01 (\*\*), *p*-value ≤ 0.001 (\*\*\*) and *p*-value ≤ 0.0001 (\*\*\*\*). **Figure 2.** (**A**) Growth of *P. brasiliensis* in the presence of cell wall disrupting agents. A total of <sup>1</sup> <sup>×</sup> <sup>10</sup><sup>6</sup> cells/mL were subjected to 25 µg/mL of iDrk1 and incubated for 24 h. Next, yeast cells were diluted, plated in solid mYPD medium containing different agents that disturb the cell wall, such as Calcofluor White (CFW), Congo Red and NaCl. Finally, cells were incubated in for seven days at 37 ◦C. Expression of cell wall morphogenesis related genes in *P. brasiliensis*. Pb18 cells were subjected to 25 µg/mL of iDrk1 for 24 h and then total RNA extraction was performed. The related genes (**B**) synthesis of chitin, such as *CHT3*, *CHS2*, *CHS3*, *CHS4* and *CHS5* and (**C**) synthesis of glucans such as *FKS*, *KRE6*, *PHR2*, *GEL3* and *AGN1* were analyzed, where *p*-value ≤ 0.01 (\*\*), *p*-value ≤ 0.001 (\*\*\*) and *p*-value ≤ 0.0001 (\*\*\*\*).

Yeasts were incubated for 24 h in the presence or absence of iDrk1. Then, labelling was performed for each fluorescent marker described above. Fluorescence analyses were obtained through flow cytometry, where 10,000 events were obtained for each sample.

≤ 0.0001 (\*\*\*\*).

Quantification was performed using the median fluorescence intensity (MFI). Figure 3A,B show a significant increase in β-glucan and chitin levels after 24 h of treatment with iDrk compared to the untreated control. On the other hand, no change in mannan levels was observed. In addition to the flow cytometry measurements, β-glucan localization was also evaluated by confocal microscopy. Yeast labelling was performed as described in the previous section using the Dectin-1-Fc probe and a secondary antibody anti-mouse IgG conjugated to Alexa 488. This assay made it possible to observe an increase in labelling in the fungus previously incubated with iDrk1 compared to the control. Most of the marking was observed on the yeast surface (Figure 3C). Fluorescence quantification (Figure 3D) was obtained from the corrected total fluorescence values (CTCF). These results indicate that PbDrk inhibition increases the β-glucan levels in *P. brasiliensis* yeast cells. *J. Fungi* **2021**, *7*, 1014 11 of 21 was observed on the yeast surface (Figure 3C). Fluorescence quantification (Figure 3D) was obtained from the corrected total fluorescence values (CTCF). These results indicate that PbDrk inhibition increases the β-glucan levels in *P. brasiliensis* yeast cells.

**Figure 3.** Quantification of *P. brasiliensis* cell wall components after incubation with iDrk1. Pb 18 cells were subjected to 25 µg/mL of iDrk1 for 24 h and then labeled with fc-Dectin-1 and Alexa 488 for dosage of β-glucan, WSA conjugated with FITC for dosage of chitin oligomers and Concanavalin A conjugated with FITC for mannan dosage. (**A**) Histograms from **Figure 3.** Quantification of *P. brasiliensis* cell wall components after incubation with iDrk1. Pb 18 cells were subjected to 25 µg/mL of iDrk1 for 24 h and then labeled with fc-Dectin-1 and Alexa 488 for dosage of β-glucan, WSA conjugated with

each cell wall component by flow cytometry, where negative control is represented as yeast that were not fluorescence labelled. (**B**) Quantification through the median fluorescence intensity (MFI) where *p*-value ≤ 0.004 (\*\*). Evaluation of βglucan exposure in *P. brasiliensis* cell wall after incubation with iDrk1. Cells of Pb18 were subjected to 25 µg/mL of iDrk1 for 24 h and then labeled with fc-Dectin-1 and Alexa 488. (**C**) Confocal microscopy analysis. (**D**) Corrected quantification of total fluorescence (CTCF) from confocal microscopy analysis where *p*-value ≤ 0.01 (\*\*), *p*-value ≤ 0.001 (\*\*\*) and *p*-value

FITC for dosage of chitin oligomers and Concanavalin A conjugated with FITC for mannan dosage. (**A**) Histograms from each cell wall component by flow cytometry, where negative control is represented as yeast that were not fluorescence labelled. (**B**) Quantification through the median fluorescence intensity (MFI) where *p*-value ≤ 0.004 (\*\*). Evaluation of β-glucan exposure in *P. brasiliensis* cell wall after incubation with iDrk1. Cells of Pb18 were subjected to 25 µg/mL of iDrk1 for 24 h and then labeled with fc-Dectin-1 and Alexa 488. (**C**) Confocal microscopy analysis. (**D**) Corrected quantification of total fluorescence (CTCF) from confocal microscopy analysis where *p*-value ≤ 0.01 (\*\*), *p*-value ≤ 0.001 (\*\*\*) and *p*-value ≤ 0.0001 (\*\*\*\*).

## *3.5. Inhibition of PbDrk1 Induces Increased Phagocytosis of P. brasiliensis and Alters Cytokine Production by Macrophages*

Fungal cell wall β-glucan is also an important pathogen-associated molecular pattern (PAMP). To avoid the innate immune response, many fungal pathogens depend on the synthesis of the cell wall α-glucan, which functions as a stealth molecule to mask the β-glucans itself or links other masking structures to the cell wall [56]. As demonstrated in the previous results, when we inhibited the PbDrk1 pathway, the Pb18 yeast underwent changes in their cell wall composition. Under normal conditions, the main components of the outermost layer of the Pb18 cell wall are α-glucan molecules [13,57]. Thus, we determined whether the increase in β-glucan would make the fungus more susceptible to recognition by cells of the immune system. To answer this question, a phagocytosis assay was performed with J774 murine macrophages. After 24 h of interaction, the supernatant was collected for subsequent cytokine dosage. In this assay, it was possible to observe that fungi treated with iDrk1 had a higher phagocytic index than the untreated control (Figure 4A). To confirm these observations, a phagocytosis assay was performed using flow cytometry [37]. The sample treatment was performed as described above, but before the cell-yeast interaction, the Pb18 yeasts were labelled with the intracellular dye CFSE. This result is similar to that observed in the previous experiment. In Figure 4B, we can see the greater signal intensity in the sample where the fungus was previously treated with iDrk1. Furthermore, we observed a reduced number of colony-forming units (CFUs) in the group treated with iDrk1 (Figure 4C). This dataset suggests that PbDrk1 inhibition of *P. brasiliensis* may have contributed to an increase in phagocytosis and in the susceptibility of fungal cells to macrophage elimination.

The quantification of the proinflammatory cytokines TNFα and IL-12p70 was evaluated by ELISA. Before the phagocytosis assay, J774 cells were primed with LPS. As a control, the supernatant from the macrophages activated only with LPS was analysed. In Figure 4D, we can see a significant increase in TNFα levels in the supernatant of cell cultures incubated with *P. brasiliensis* yeasts previously exposed to iDrk1. The same result was observed for the cytokine IL-12p70 (Figure 4E).

#### *3.6. Regulation of cAMP-PKA and Glycogen Accumulation in P. brasiliensis*

Perception of the environment is fundamental for fungal survival in the host. The cyclic AMP-dependent protein kinase A pathway (cAMP-PKA) is highly conserved and is involved in several biological processes, both in human pathogenic and phytopathogenic fungi [58]. In addition, it also contributes to gene expression regulation and cell wall remodeling [59]. Through stimuli external to the cell, the enzyme adenylate cyclase converts ATP to adenosine 30 ,50 -cyclic monophosphate (cAMP), an important secondary messenger that binds to the catalytic subunit of protein kinase A (PKA). This generates a conformational change that releases PKA catalytic subunits and activates transcription factors and other signalling pathways involved in cell wall integrity, stress response and virulence [58,60,61]. Thus, Pb18 yeasts treated or not with iDrk1 for 24 h were lysed, and the protein extract was used to quantify the PKA activity and cAMP dosage. As shown in Figure 5A, it was possible to verify a reduction in PKA activity after exposure to iDrk1. As a positive control for the reaction, *P. brasiliensis* was subjected to oxidative stress with 300 mM hydrogen peroxide (H2O2) for 30 min. This condition is known to generate an

increase in PKA activity [62]. It was also possible to observe no significant difference in the dosage of cAMP levels between the treated and control samples (Figure 5B).

*J. Fungi* **2021**, *7*, 1014 13 of 21

**Figure 4.** Phagocytosis assay. 2.5 × 105 J774 cells (MΦ) were plated in RPMI medium supplemented with 10% FBS. Then the cells were primed for 30 min with LPS 100 ng/mL. In parallel, *P. brasiliensis*  yeasts were subjected to 25 µg/mL of iDrk1 for 24 h. The interaction was carried out in a 2:1 ratio (yeast:macrophages) for 24 h at 37 °C and 5% CO2. After this period, the cells were stained and visualized under an optical microscope to count the internalized yeasts. (**A**) Phagocytic index was calculated, where *p*-value ≤ 0.005 (\*\*). (**B**) Phagocytosis assay performed by labeling *P. brasiliensis*  yeasts with CFSE prior to interaction with macrophages. Fluorescence quantification was performed by flow cytometry and relative quantification was obtained through the median intensity (MFI) where *p*-value ≤ 0.005 (\*\*). (**C**) CFU determination from yeasts recovered from macrophages, where *p*-value ≤ 0.0001 (\*\*\*\*). Cytokine assay from J774 cell supernatant after interaction with *P. brasiliensis*  submitted or not with iDrk1. Macrophages were plated in RPMI medium supplemented with 10% FCS. Then, cells were primed for 30 min with LPS 100 ng/mL. In parallel, *P. brasiliensis* yeasts were subjected to 25 µg/mL of iDrk1 for 24 h. The interaction was carried out in a 2:1 ratio (yeast:macrophages) for 24 h at 37 °C and 5% CO2. After this period, the supernatant was collected and the cytokines (**D**) TNFα, (**E**) IL-12p70 were measured, where *p*-value ≤ 0.05 (\*), *p*-value ≤ 0.01 (\*\*). **Figure 4.** Phagocytosis assay. 2.5 <sup>×</sup> <sup>10</sup><sup>5</sup> J774 cells (MΦ) were plated in RPMI medium supplemented with 10% FBS. Then the cells were primed for 30 min with LPS 100 ng/mL. In parallel, *P. brasiliensis* yeasts were subjected to 25 µg/mL of iDrk1 for 24 h. The interaction was carried out in a 2:1 ratio (yeast:macrophages) for 24 h at 37 ◦C and 5% CO<sup>2</sup> . After this period, the cells were stained and visualized under an optical microscope to count the internalized yeasts. (**A**) Phagocytic index was calculated, where *p*-value ≤ 0.005 (\*\*). (**B**) Phagocytosis assay performed by labeling *P. brasiliensis* yeasts with CFSE prior to interaction with macrophages. Fluorescence quantification was performed by flow cytometry and relative quantification was obtained through the median intensity (MFI) where *p*-value ≤ 0.005 (\*\*). (**C**) CFU determination from yeasts recovered from macrophages, where *p*-value ≤ 0.0001 (\*\*\*\*). Cytokine assay from J774 cell supernatant after interaction with *P. brasiliensis* submitted or not with iDrk1. Macrophages were plated in RPMI medium supplemented with 10% FCS. Then, cells were primed for 30 min with LPS 100 ng/mL. In parallel, *P. brasiliensis* yeasts were subjected to 25 µg/mL of iDrk1 for 24 h. The interaction was carried out in a 2:1 ratio (yeast:macrophages) for 24 h at 37 ◦C and 5% CO<sup>2</sup> . After this period, the supernatant was collected and the cytokines (**D**) TNFα, (**E**) IL-12p70 were measured, where *p*-value ≤ 0.05 (\*), *p*-value ≤ 0.01 (\*\*).

Perception of the environment is fundamental for fungal survival in the host. The cyclic AMP-dependent protein kinase A pathway (cAMP-PKA) is highly conserved and is involved in several biological processes, both in human pathogenic and phytopathogenic fungi [58]. In addition, it also contributes to gene expression regulation and cell wall remodeling [59]. Through stimuli external to the cell, the enzyme adenylate cyclase converts ATP to adenosine 3′,5′-cyclic monophosphate (cAMP), an important secondary mes-

formational change that releases PKA catalytic subunits and activates transcription factors and other signalling pathways involved in cell wall integrity, stress response and virulence [58,60,61]. Thus, Pb18 yeasts treated or not with iDrk1 for 24 h were lysed, and the protein extract was used to quantify the PKA activity and cAMP dosage. As shown in

*3.6. Regulation of cAMP-PKA and Glycogen Accumulation in P. brasiliensis* 

glycogen accumulation [65,66].

**Figure 5.** PKA activity and cAMP quantification in *P. brasiliensis* after incubation with iDrk1 (**A**) Quantification of PKA activity and (**B**) dosage of *P. brasiliensis* cAMP after 24 h incubation with 25 µg/mL of iDrk1, where *p*-value ≤ 0.001 (\*\*\*). (**C**) After 24 and 48 h of yeast incubation in the presence of the inhibitor, the cells were stained with a 0.2% iodine and **Figure 5.** PKA activity and cAMP quantification in *P. brasiliensis* after incubation with iDrk1 (**A**) Quantification of PKA activity and (**B**) dosage of *P. brasiliensis* cAMP after 24 h incubation with 25 µg/mL of iDrk1, where *p*-value ≤ 0.001 (\*\*\*). (**C**) After 24 and 48 h of yeast incubation in the presence of the inhibitor, the cells were stained with a 0.2% iodine and 0.4% potassium iodide solution to assess glycogen accumulation.

**4. Discussion**  Currently, the number of antifungal substances available for the treatment of phytopathogens is approximately nine times greater than those available for the treatment of mycoses in mammals [67]. In this scenario, it is important to emphasize the need for studies that unravel the mechanisms of fungal pathogenicity. With a better understanding of these pathways and the discovery of new targets, it will be possible to develop new drugs with antifungal potential. Thus, this study aimed to understand better the role of a histidine kinase (PbDrk1), an important regulator in the dimorphic switch and morphogenesis of the cell wall of *P. brasiliensis*. We evaluated pharmacological inhibitors as a strategy to elucidate the role of PbDrk1. The inhibitors iprodione and fludioxonil (both group III histidine kinase inhibi-The cell stress response is orchestrated by connecting several pathways that converge to promote cell survival. As previously described, the cell wall integrity pathway also activates responses that bind to cAMP-PKA. Activation of PKA regulates cellular functions related to glycogen metabolism [63], and it antagonizes its intracellular accumulation [64]. Thus, Pb18 yeasts incubated in the presence or absence of iDrk1 for 24 and 48 h were stained with iodine solution and photographed (Figure 5C). The greater the accumulation of intracellular glycogen, the darker the sample becomes [33]. In Figure 5C, there was an accumulation of intracellular glycogen in relation to the control. Combined with the gene expression analysis and quantification of the cell wall components, this result suggests that iDrk1 may modulate the cell wall component synthesis, generating a cellular response that leads to a decrease in PKA activity. This decreased activity may lead to intracellular glycogen accumulation [65,66].

**.** 

Figure 5A, it was possible to verify a reduction in PKA activity after exposure to iDrk1. As a positive control for the reaction, *P. brasiliensis* was subjected to oxidative stress with 300 mM hydrogen peroxide (H2O2) for 30 min. This condition is known to generate an increase in PKA activity [62]. It was also possible to observe no significant difference in

The cell stress response is orchestrated by connecting several pathways that converge to promote cell survival. As previously described, the cell wall integrity pathway also activates responses that bind to cAMP-PKA. Activation of PKA regulates cellular functions related to glycogen metabolism [63], and it antagonizes its intracellular accumulation [64]. Thus, Pb18 yeasts incubated in the presence or absence of iDrk1 for 24 and 48 h were stained with iodine solution and photographed (Figure 5C). The greater the accumulation of intracellular glycogen, the darker the sample becomes [33]. In Figure 5C, there was an accumulation of intracellular glycogen in relation to the control. Combined with the gene expression analysis and quantification of the cell wall components, this result suggests that iDrk1 may modulate the cell wall component synthesis, generating a cellular response that leads to a decrease in PKA activity. This decreased activity may lead to intracellular

the dosage of cAMP levels between the treated and control samples (Figure 5B).

#### tors) are substances widely used in agriculture to combat phytopathogens [68]. Molecular **4. Discussion**

0.4% potassium iodide solution to assess glycogen accumulation.

biology and cell signalling studies have shown that these molecules act specifically on group III histidine kinases through interactions with HAMP domains, which are present only in this group of kinases [39,69]. In *Saccharomyces cerevisiae*, group III histidine kinases are absent. When an orthologous gene to histidine kinase group III of *Neurospora crassa* is introduced, *S. cerevisiae* becomes sensitive to pyrrolnitrine [68,70]. Other pathogenic fungi Currently, the number of antifungal substances available for the treatment of phytopathogens is approximately nine times greater than those available for the treatment of mycoses in mammals [67]. In this scenario, it is important to emphasize the need for studies that unravel the mechanisms of fungal pathogenicity. With a better understanding of these pathways and the discovery of new targets, it will be possible to develop new drugs with antifungal potential. Thus, this study aimed to understand better the role of a histidine kinase (PbDrk1), an important regulator in the dimorphic switch and morphogenesis of the cell wall of *P. brasiliensis*.

We evaluated pharmacological inhibitors as a strategy to elucidate the role of PbDrk1. The inhibitors iprodione and fludioxonil (both group III histidine kinase inhibitors) are substances widely used in agriculture to combat phytopathogens [68]. Molecular biology and cell signalling studies have shown that these molecules act specifically on group III histidine kinases through interactions with HAMP domains, which are present only in this group of kinases [39,69]. In *Saccharomyces cerevisiae*, group III histidine kinases are absent. When an orthologous gene to histidine kinase group III of *Neurospora crassa* is introduced, *S. cerevisiae* becomes sensitive to pyrrolnitrine [68,70]. Other pathogenic fungi with group III histidine kinases, such as *B. dermatitidis* and *Candida albicans*, were submitted to this inhibitor and presented various sensitivities [17]. For Pb18 yeast cells, fludioxonil was used at 100 µg/mL, showing no reduced cell viability. Thus, the concentration established in the other tests at 25 µg/mL was selected since it is an intermediate concentration and was already established in *B. dermatitidis* [17].

The fungal cell wall plays a fundamental role in the host-parasite interaction since its composition can influence the immune response [13]. In *Paracoccidioides* spp., during

the dimorphic transition, the cell wall morphogenesis machinery is activated to remodel its components. In *H. capsulatum* and *B. dermatitidis,* it was shown that *DRK1* acts as a regulator of dimorphism and virulence. A study carried out in *B. dermatitidis* silenced the *DRK1* gene and demonstrated a blockade of the dimorphic transition from mycelium to yeast at 37 ◦C. Furthermore, this silencing impaired the expression of *BAD1* (a virulence gene activated during transition) [11]. In *H. capsulatum*, Drk1 regulates genes specifically expressed in the yeast phase, such as *CBP1*, *YPS-3* and *AGS1* [11]. In *C. albicans*, the deletion of *NIK1*, a group III histidine kinase, makes yeast incapable of transitioning from yeast to hyphae, consequently making it less virulent [71]. In *Penicillium marneffei*, the deletion of the *DRKA* and *SLNA* genes was also essential for the dimorphic transition during macrophage infection and conidial germination, respectively [44].

The ability of *Paracoccidioides* spp. to cause PCM depends on its dimorphic transition and establishment in the host, either by the resistance and evasion of the immune response machinery or by its ability to be a facultative intracellular parasite [8]. The use of iDrk1 (fludioxonil) in *P. brasiliensis* cultivated in the form of mycelium prevented it from performing the dimorphic transition efficiently, confirming previous findings [20]. A study carried out in *Penicillium marneffei* showed the importance of *DRK1* in cell wall morphogenesis. Strains deleted for this gene were not able to grow in a medium supplemented with cell wall stressors (Congo red) and osmotic stress agents (NaCl and sorbitol) [44]. In that same study, *DRK1* mutant hyphal growth and conidial germination were affected, and transmission electron microscopy images showed significant thickening of the cell wall [44]. These data support the results obtained in Pb18. In this work, yeasts submitted to iDrk1 and later inoculated in culture medium supplemented with cell wall stressors had severely impaired growth.

In *Paracoccidioides* spp. the cell wall chitin content represents a significant fraction of the cell dry weight [49]. The *CHT3*, *CHS2,* and *CHS3* genes showed significant increases compared with the control that was not treated with iDrk1. In *C. albicans*, it was observed that overexpression of *CHT3* induces greater sensitivity to cell wall stressors, and the *CHS2* gene acts as an important regulator [72,73]. In *P. brasiliensis*, it is known that the *CHS3* gene is mainly expressed in the yeast phase [49] and is related to cell growth [74]. These results suggest that PbDrk1 inactivity may modulate the expression of some genes involved in chitin synthesis, leading to cell wall instability.

In addition to chitin, one of the main elements that comprise the cell wall of *Paracoccidioides* spp. are glucan molecules. As already mentioned, the predominance of β-glucan during the mycelial phase is reverted to α-glucan in the yeast phase [2]. This mechanism is seen as one strategy to evade the immune system. The level of α-glucan in the cell wall can be related to the degree of virulence [14]. Among the genes analysed, *AGN1* was characterized as phase-specific in *P. brasiliensis* yeasts, regulating the synthesis of α-(1,3)-glucan [54]. Our data showed that PbDrk1 does not participate in the modulation of this gene. On the other hand, genes involved in the synthesis and maintenance of β-(1,3)-glucan and β-(1,6)-glucan chains, such as *FKS1*, *KRE6*, *PHR2*, and *GEL3*, showed significant increases after exposure to iDrk1. These results indicate that PbDrk1 regulates the expression of cell wall synthesis genes directly related to the dimorphic transition. The expression data were confirmed by evaluating the levels of β-glucan and chitin in Pb18 treated with a Drk1 inhibitor. The glucan modulation transition is essential for masking β-glucan molecules, as the host's immune system directly recognizes these molecules by phagocytic cells.

In the host, the fungus is initially recognized by PRRs and PAMPs [75]. Among the receptors that may be involved, we highlight the Dectin-1 receptor that specifically binds to β-glucan [76]. Chitin molecules are also recognized by the immune system via TLR-2, inducing the production of cytokines and the recruitment of phagocytic cells by the recognition by Dectin-1 [77]. The outermost region of the *P. brasiliensis* cell wall comprises a thick layer of mannan [2]. This layer is believed to protect fungal cells from immunological recognition, hiding the main immunogenic molecules [78]. Confocal microscopy assays

indicated an increase in β-glucan labelling. These images showed that yeasts treated with iDrk1 showed greater fluorescence intensity when labelled with the Dectin-1 receptor. Thus, the increase in β-glucan and increased expression and synthesis of chitin in cells treated with iDrk1 indicate that this pathway may be related to virulence and fungal cell wall architecture.

In parallel, the cAMP-PKA pathway also plays a role in cell wall remodeling. In *Cryptococcus neoformans*, PKA is involved in the phosphorylation of components that, when translocated to the nucleus, regulate the expression of genes involved in cell wall integrity [59]. In fungi, this pathway is related to cell growth, differentiation, stress response, pathogenicity, and cell wall integrity, among others [61]. In *C. neoformans,* it was shown that the deletion of several genes involved in cell wall integrity led to impairment of the cAMP-PKA pathway [79]. In *C. albicans*, this pathway plays a central regulator of its morphological transition and, consequently, pathogenicity [80]. In *P. lutzii,* it was reported that the specific inhibition of PKA impedes dimorphic transition [81]. Recently, Garcia et al.; (2017) showed a new insight into the signalling pathways involved in the regulation of cell wall integrity. Alterations of the β-1,3-glucan network in the fungal cell wall induced the activation of the CWI pathway and in parallel inhibited the PKA signalling [33]. Thus, when submitting Pb18 yeasts to iDrk1, we observed a reduction in PKA activity, supporting literature data on the PKA influence on cell wall gene expression and glycogen accumulation [33,59] and a possible correlation with the PbDrk1 pathway.

On the other hand, there was no significant difference in cAMP levels in the presence or absence of iDrk1. Cyclic AMP is a secondary messenger whose intracellular levels are regulated by adenylate cyclase and the phosphodiesterase enzyme balance [65]. Both are regulated by other signalling pathways and not exclusively via PKA [82]. Thus, it is possible to infer that PbDrk1 modulates PKA activity but does not correlate with cAMP levels. In this case, the maintenance of cAMP levels can be regulated by other pathways, such as calcineurin, MAP kinases, and G protein subunits [80,83]. Finally, we can infer that PbDrk1 modulates intracellular levels of intracellular glycogen via PKA. In *Aspergillus fumigatus*, deletion of the PKA catalytic subunit led to an increase in intracellular glycogen levels [65]. Furthermore, in *S. cerevisiae*, it was found that glycogen is present in two fractions, one of them in the cell cytosol in its soluble form and the other associated with cell wall components, covalently linked specifically to β-(1,3)-glucan and β-(1,6)-glucan [84]. This evidence complements the observed results of increased β-glucan exposure on the surface of *P. brasiliensis* after exposure to iDrk1. Together, the results obtained thus far point to the participation of PbDrk1 in the *P. brasiliensis* cell wall modulation.

During the immune response in a fungal infection, the action of pro- and antiinflammatory cytokines is essential to determine disease progression and/or pathogen clearance [85]. Host resistance to infection by *Paracoccidioides* spp. is associated with a Th1 response, which induces macrophage activation and actively controls fungal dissemination [86]. The individual's susceptibility to the disease is associated with a Th2 response [87]. It is known that the participation of the Dectin-1 receptor together with TLR in the recognition pathway of *Paracoccidioides* spp. triggers the production of several proinflammatory cytokines [88], such as TNFα. This molecule is produced by cells involved in the immune system, including activated macrophages and regulatory T cells, acting as central mediators in inflammation and the regulation of the immune response [89]. It is possible to observe the increased production of this cytokine by macrophages that phagocytosed yeasts exposed to iDrk1, indicating a more exacerbated proinflammatory response when compared to the control. Furthermore, a significant increase in the expression of the cytokine IL12p70 was also observed, produced by monocytes, macrophages, and dendritic cells, and it is an important element in the activation of the Th1 response [90].

#### **5. Conclusions**

The data obtained during this work will contribute to a better understanding of a regulatory pathway that has not yet been explored in this model. The PbDrk1 protein has

already been shown to be a key element in the dimorphic transition pathway [20]. It is now possible to state that it is a likely virulence factor in regulating cell wall genes. The activity of PbDrk1 negatively modulates the synthesis of molecules such as chitin and β-glucans, contributing to its masking and favouring the pathogenicity of *P. brasiliensis*. Histidine kinases are proteins absent in mammals, and the inhibition of their activity makes fungal cells susceptible to immune system cells. In other fungal models, the loss of this gene represents a decrease in virulence. Finally, the study of PbDrk1 and associated pathways would enable a different approach to the development of new antifungal drugs.

**Author Contributions:** Conceptualization, W.L.B. and M.V.N.; methodology, M.V.N., Y.N.d.B., W.D.S., A.F.A.C., K.S.F., G.P.J. and P.X.; software, M.V.N., Y.N.d.B., W.D.S.; validation, M.V.N. and Y.N.d.B.; formal analysis, M.V.N.; writing—review and editing, W.L.B. and M.V.N.; supervision, W.L.B.; project administration, W.L.B. and M.V.N.; funding acquisition, W.L.B. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP), grant number 2017/04592-0 and Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) grant number 141726/2017-2; 311008/2020-8 and Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES).

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Acknowledgments:** We are thankful to Sandro Rogério de Almeida (Universidade de São Paulo— USP, Brazil) for technical assistance and material donation to ELISA and phagocytosis experiments. We are thankful for the support of Núcleo de Microscopia Confocal Unifesp Diadema for the confocal microscopy experiments.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

