The human fungal pathogen, Cryptococcus neoformans, dramatically alters its cell wall, both in size and composition, upon entering the host. This cell wall remodeling is essential for host immune avoidance by this pathogen. In a genetic screen for mutants with changes in their cell wall, we identified a novel protein, Mar1, that controls cell wall organization and immune evasion. Through phenotypic studies of a loss-of-function strain, we have demonstrated that the mar1Δ mutant has an aberrant cell surface and a defect in polysaccharide capsule attachment, resulting in attenuated virulence. Furthermore, the mar1Δ mutant displays increased staining for exposed cell wall chitin and chitosan when the cells are grown in host-like tissue culture conditions. However, HPLC analysis of whole cell walls and RT-PCR analysis of cell wall synthase genes demonstrated that this increased chitin exposure is likely due to decreased levels of glucans and mannans in the outer cell wall layers. We observed that the Mar1 protein differentially localizes to cellular membranes in a condition dependent manner, and we have further shown that the mar1Δ mutant displays defects in intracellular trafficking, resulting in a mislocalization of the β-glucan synthase catalytic subunit, Fks1. These cell surface changes influence the host-pathogen interaction, resulting in increased macrophage activation to microbial challenge in vitro. We established that several host innate immune signaling proteins are required for the observed macrophage activation, including the Card9 and MyD88 adaptor proteins, as well as the Dectin-1 and TLR2 pattern recognition receptors. These studies explore novel mechanisms by which a microbial pathogen regulates its cell surface in response to the host, as well as how dysregulation of this adaptive response leads to defective immune avoidance.
Disease causing microorganisms have adapted many strategies to avoid host immune detection in order to facilitate their survival. Pathogenic fungi alter their cell surface in order to mask immune stimulatory epitopes in their cell wall. We have identified a novel protein involved in this host-induced cell wall remodeling process in Cryptococcus neoformans. We have created a mutant strain with a targeted deletion of this gene and demonstrated that this protein is involved in intracellular trafficking of the β-(1,3)-glucan synthase enzyme and is differentially localized in response to host-like conditions. In the absence of this protein, the cell wall has decreased levels of β-glucan and increased exposure of chitooligomers. Importantly, improper cell wall maintenance by this mutant leads to a hyper-immunostimulatory fungal cell surface that induces increased macrophage activation. We show that this activation is dependent on several pattern recognition receptors previously demonstrated to be important for fungal pathogenesis.
Citation: Esher SK, Ost KS, Kohlbrenner MA, Pianalto KM, Telzrow CL, Campuzano A, et al. (2018) Defects in intracellular trafficking of fungal cell wall synthases lead to aberrant host immune recognition. PLoS Pathog 14(6): e1007126. https://doi.org/10.1371/journal.ppat.1007126
Editor: Robin Charles May, University of Birmingham, UNITED KINGDOM
Received: January 20, 2018; Accepted: May 29, 2018; Published: June 4, 2018
Copyright: © 2018 Esher et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: These experiments were supported by a National Institutes of Health grant awarded to JAA and FLW, Jr. (R01 AI074677, https://grants.nih.gov/grants/oer.html). CM and colleagues Jeanette Wagener, Louise Walker, Neil Gow were supported by the Wellcome Trust Strategic Award in Medical Mycology and Fungal Immunology (097377, https://wellcome.ac.uk), Wellcome Trust Senior Investigator Award (101873) and the MRC Centre for Medical Mycology (MR/N006364/1, https://www.abdn.ac.uk/cmm/). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
The microbial surface is the first point of contact for interactions with the innate immune system, representing the site at which an infected host might recognize a microbe as a potential pathogen. This recognition is achieved through host pattern recognition receptors (PRRs) that distinguish specific pathogen-associated molecular patterns (PAMPs) on microbial surfaces, directing downstream signaling events that ultimately lead to the initiation of an immune response. The fungal cell wall is a dynamic structure composed of a complex matrix of polysaccharides including α- and β-glucans, mannoproteins (mannans), and chitin/chitosan. These fungal specific components have been shown by many groups to be recognized by host PRRs including Toll-like receptors (TLRs) and C-type lectin receptors (CLRs) [1,2]. Several fungi have developed strategies to mask their surfaces from immune detection. Examples include Histoplasma capsulatum cell wall α-(1,3)-glucan and Aspergillus fumigatus conidial RodA hydrophobin, which both serve to block exposure of the more immunogenic β-glucan molecule [3,4]. The fungal cell surface is also responsive to different environments, including various micro-environments within the infected host. For example, Candida albicans differentially exposes β-glucan in response to diverse host niches, drug treatments, and growth conditions, resulting in varying degrees of Dectin-1-mediated host responses [5,6].
The opportunistic human fungal pathogen Cryptococcus neoformans continues to be a significant health threat for immune compromised populations, particularly those with HIV/AIDS, among whom it causes over 175,000 deaths per year . This ubiquitous fungus colonizes the lungs after inhalation from the environment. It can then disseminate to the central nervous system in immunocompromised individuals, where it causes life-threatening meningoencephalitis . C. neoformans has developed several adaptations to avoid immune detection and to direct the host immune response in its favor. The polysaccharide capsule on the cell surface shields immunostimulatory cell wall components from host recognition. Additionally, secreted capsular material actively represses various immune responses . C. neoformans cells can also grow to massive sizes, forming so-called titan cells in the setting of infection. These giant and hyper-encapsulated cells are unable to be engulfed by host immune cells, but instead they drive a non-protective immune response to C. neoformans leading to pathogen persistence [10–12].
Both capsule production and titan cell formation are induced in the host environment and involve significant cell wall remodeling. Previous work has shown that capsule polysaccharide likely attaches to the cell surface through interaction with α-(1,3)-glucan in the cell wall . Titan cell walls are also thicker and more chitin-rich then normal sized cells . In addition to these macromolecular cell surface changes, our lab has shown that C. neoformans actively remodels its cell wall in response to host pH signals in order to avoid immune detection [14–16]. We have recently demonstrated that aberrant chitooligomer exposure leads to a detrimental immune response . Although few investigators have performed detailed cell wall analyses in C. neoformans, this structure appears to contain significantly more chitin and chitosan than that of other pathogenic species . There is increasing evidence to support the concept of chitin as an immune modulatory molecule, and recent studies have highlighted the complexity of chitin recognition, indicating that the source and size of chitin molecules can differentially direct immune responses [18–20].
C. neoformans cell wall regulation, particularly in response to the host environment, remains incompletely defined. Our lab has previously identified cell surface properties that drive immune detection and through a previously published screen for cell wall regulators, we have identified a novel protein involved in this process [14,21,22]. In this study we explore the mechanism by which this protein, Mar1, regulates cell wall remodeling, and the implications of an aberrant cell wall architecture on host immune detection. We report that in the absence of Mar1, cells display a defect in capsule attachment, as well as increased exposure of cell wall chitin and chitosan. This concurs with our observation of decreased levels of glucans and mannans in the cell wall, decreased expression of α- and β-glucan synthases, and mislocalization of the β-glucan synthase, Fks1. While canonical secretion is intact in the mar1Δ mutant strain, general intracellular trafficking appears altered. We also report the localization of the Mar1 protein itself to cellular membranes and demonstrate differential localization in host-like tissue culture conditions. The implications of this cell wall remodeling defect include increased recognition and activation by innate immune cells and attenuated virulence. We further show that this innate immune recognition is dependent on the Card9 and MyD88 adaptor proteins and the cell surface receptors Dectin-1 and TLR2. In addition to highlighting intracellular processes involved in cell wall remodeling, these studies underscore the importance of proper cell wall regulation in host immune detection and broaden our understanding of the recognition of individual fungal cell wall components.
Identification of a novel cell wall regulatory protein in Cryptococcus neoformans
The cell wall is the interface between microbial pathogens and host immune cells. To identify genes required for proper fungal cell wall homeostasis, we performed a random mutagenesis screen in the human fungal pathogen Cryptococcus neoformans using Agrobacterium tumefaciens-mediated transformation (AMT). Mutagenized strains were screened for phenotypes corresponding to cell wall changes, including growth impairment and dry colony morphology in alkaline conditions (pH 8) and sensitivity to elevated salt concentrations (1.5 M NaCl). We hypothesized that a subset of these cell wall mutants would also display alterations in the host-pathogen interaction. Preliminary results from this screen were previously reported .
From this screen, we identified a mutant displaying dry colony morphology and slight sensitivity to alkaline pH as well as decreased growth on elevated salt concentrations. This strain had a mutation in a previously uncharacterized gene (CNAG_06695), which we have named MAR1 (macrophage activating regulator of cell wall-1). The encoded protein is predicted to contain two transmembrane domains, which comprise a larger domain of unknown function (DUF4112) (Fig 1A). However, Mar1 has no annotated predicted functions based on its sequence, and it shares no significant sequence homology with proteins from other species, including other basidiomycetes, with the exception of highly related Cryptococcus species (C. deneoformans, C. deuterogattii, and C. gattii). We confirmed the phenotypes of this mutant by independently disrupting the entire MAR1 gene in the wild type background (Fig 1B). We also complemented all phenotypes by reintroduction of the wild type allele into the mar1Δ mutant.
(A) Schematic of the Mar1 protein domains. DUF, domain of unknown function; TM, transmembrane. (B) The mar1ΔT-DNA insertional mutant and a mar1Δ full deletion mutant are dry on pH 8 and sensitive to 1.5 M NaCl. Serial dilutions of the indicated strains were spotted onto YPD, YPD with 150 mM HEPES at pH 8, or YPD with 1.5 M NaCl. (C) The mar1Δ strain shares distinct and overlapping sensitivities to cell wall stressors with other cell wall integrity pathway mutants. Strains were serially diluted and spotted onto YPD with the addition of the indicated cell wall stressors or at the indicated temperatures. Melanin was assessed by the production of brown pigment on Niger Seed agar at 30°C. CFW, calcofluor white; SDS, sodium dodecyl sulfate.
We assessed the sensitivity of the mar1Δ mutant to common cell wall stressors. These agents included calcofluor white (CFW, binds and blocks chitin assembly), Congo red (inhibits assembly of cell wall polymers, especially chitin), caffeine (affects signal transduction and general cell wall integrity), and SDS (cell membrane stressor) [23–26]. When incubated in the presence of cell wall stressors, growth of the mar1Δ strain was severely inhibited by Congo red (0.5%) and SDS (0.06%) compared to the WT strain (Fig 1C). In contrast, the mar1Δ mutant only displayed a slight decrease in colony size on CFW (1 mg/ml), and growth was comparable to WT when incubated in the presence of caffeine (1 mg/ml).
Given the enhanced susceptibilities of the mar1Δ mutant to cell wall perturbing agents, we considered that the Mar1 protein might be involved in other cell-signaling pathways that regulate or respond to defects in cell wall integrity, including (1) the Rim/alkaline response pathway (2) the PKA/cAMP pathway, (3) the PKC/cell wall integrity (CWI) pathway, (4) the calcineurin pathway, and (5) the high-osmolality-glycerol (HOG) pathway. Therefore, we compared the mar1Δ mutant to strains with mutations in these other pathways (representative mutants used for each pathway: (1) rim101Δ; (2) pka1Δ; (3) mpk1Δ; (4) cna1Δ; (5) hog1Δ) to specifically test patterns of susceptibility to common cell wall stressors (Fig 1C). While sensitivity to Congo red and SDS is observed in mutants of the PKC/CWI and calcineurin pathways, these strains are also sensitive to caffeine. In a manner distinct from the SDS susceptibility of the mar1Δ strain, Rim and PKA/cAMP pathway mutants grew more robustly on SDS. Hog pathway strains were exclusively sensitive to SDS and no other stressors tested.
Additionally, we tested these strains for phenotypes associated with virulence including melanin production and the ability to grow at high temperature. The mar1Δ mutant displayed a modest growth defect when incubated at 37°C, however melanin production was comparable to WT (Fig 1C). These phenotypes contrasted sharply with calcineurin pathway mutants, which display defective thermotolerance, and PKA pathway mutants, that have defective melanin production [27,28]. Due to the modest temperature sensitivity of the mar1Δ strain, we also tested the effect of the Hsp90 inhibitor, radicicol, on mar1Δ growth at 30°C and 37°C. We observed identical MICs for WT and mar1Δ cells (12.5 μM at 30°C and 1.56 μM at 37°C). Together these data indicate that MAR1 is required for normal cell wall integrity under certain cell wall stress conditions. However, this particular combination of sensitivities does not precisely mimic that of mutants in any of the previously studied cell wall responsive pathways.
MAR1 is required for capsule attachment
The C. neoformans cell wall serves as the site of attachment for the polysaccharides that comprise the cell surface capsule . Therefore, some C. neoformans strains with defects in cell wall organization and structure also display defective encapsulation. We incubated the mar1Δ strain in capsule-inducing tissue culture medium and assessed capsule microscopically by India ink staining. Compared to WT, the mar1Δ mutant displayed a marked reduction in surface capsule (Fig 2A). Capsular polysaccharide is synthesized in the cytoplasm, and then secreted, where it binds to the cell surface . To differentiate between a capsule biosynthesis and capsule attachment defect, we assayed the relative amount of secreted capsule in the culture supernatant using a previously described immuno-blotting technique [21,30]. In brief, we used the mAb18B7 monoclonal antibody directed against the main capsule component GXM (glucuronoxylomannan) to probe for secreted capsule polysaccharide. By this method, we observed WT-levels of this capsular polysaccharide secreted by the mar1Δ mutant (Fig 2B). However, the electrophoretic mobility of this polysaccharide appears to differ slightly from the WT and complemented strains. These results indicate that mar1Δ has no defect in total capsule polysaccharide production, but that the defect in surface encapsulation is likely due to a defect in capsule attachment, and perhaps polysaccharide composition/structure. This pattern of altered capsule attachment is distinct from PKA pathway mutants that display impaired capsule synthesis ; however, capsule attachment defects have been observed in other strains with cell wall defects [13,21].
(A) The mar1Δ strain has a capsule defect. Cells were incubated in capsule-inducing conditions (CO2-independent tissue culture (TC) medium, 37°C) with shaking for 72 hours. Capsule was assessed by India ink counterstaining, followed by imaging. (B) The mar1Δ strain sheds capsule comparable to WT. Shed capsule polysaccharide was measured by blotting of culture supernatant, using an anti-GXM antibody to probe for capsule as described previously . (C) The mar1Δ mutant displays increased staining for exposed chitin and chitooligomers in tissue culture medium. Cells were incubated for 16–18 hours at 30°C in rich medium (YPD) or 37°C in tissue culture medium (TC), followed by staining with FITC-conjugated wheat germ agglutinin (WGA) for exposed chitin/chitooligomers, or calcofluor white (CFW) for total chitin. Stained cells were imaged by fluorescent microscopy with the appropriate filters. (D) Average fluorescence of at least 100 individual cells was measured using ImageJ/Fiji software. ****, p < 0.0001 as determined by two-way ANOVA with Tukey’s multiple comparisons test. (E) The mar1Δ mutant displays increased chitosan staining. Cells were incubated for 16–18 hours at 30°C (YPD) or 37°C (TC), followed by staining with eosin Y (EY) for chitosan. Stained cells were imaged by fluorescent microscopy. (F) The mar1Δ cell wall does not have increased total chitin or chitosan. Cells were incubated for 16–18 hours at 37°C in TC medium, followed by cell wall isolation. Chitin and chitosan levels were quantified using a modified 3-methyl-2-benzothiazolinone hydrazine hydrochloride (MBTH) colorimetric assay as described previously . Data represent means of 3 independent cell wall isolations (n = 3 for each strain). Ns, not significant as determined by two-way ANOVA with Sidak’s multiple comparisons test.
The mar1Δ mutant has an aberrant cell surface
To more fully explore the cell wall changes of the mar1Δ mutant, we performed microscopy with stains and antibodies specific for various cell wall components. After the mar1Δ mutant was incubated in host-like tissue-culture medium (TC), we observed a striking increase in its staining by FITC-conjugated wheat germ agglutinin (WGA), a lectin that recognizes exposed chitin and chitooligomers (Fig 2C) [16,31]. This pattern of increased WGA staining does not occur when the cells are incubated in rich medium (YPD), indicating this chitooligomer-exposure phenotype is induced by host-like TC conditions (Fig 2C and 2D). These initial observations were quantified, demonstrating a significant increase in the average fluorescence of WGA-stained mar1Δ cells incubated in TC medium compared to WT cells or mar1Δ cells incubated in YPD (Fig 2D).
Whereas chitooligomer exposure is well approximated by quantifying cell surface binding of the large WGA molecule, total cell wall chitooligomer content is better assessed using calcofluor white (CFW), a smaller fluorescent molecule that more easily penetrates into the cell surface and binds chitin and other chitooligomers. In contrast to the marked shift in WGA staining, we only observed a slight increase in CFW staining of mar1Δ cells compared to WT, and only when the cells were incubated in TC medium (Fig 2C and 2D).
We also measured levels of chitosan, the de-acetylated form of chitin, by Eosin Y (EY) staining (Fig 2E) . Similar to staining for chitin, we did not observe a major change in fluorescence when cells were incubated in YPD (Fig 2E); however, we observed enhanced staining of mar1Δ cells incubated in TC medium compared to WT cells. Together these cell wall staining results show that in host-like tissue culture conditions the mar1Δ strain has increased exposure of chitooligomers, including both chitin and chitosan.
The mar1Δ mutant cell wall changes in tissue culture medium are induced by pH and glucose deprivation
To better understand how host-like tissue culture conditions induce cell wall changes in the mar1Δ mutant, we sought to better define what component of this condition is responsible. To determine if the mar1Δ cell wall changes were dependent on temperature, we incubated cells in YPD or TC media at 30°C and 37°C, and measured staining of exposed chitooligomers by WGA (Fig 3A and 3B, S1A Fig). We observed a small, but not significant, increase in staining of cells incubated in YPD at 37°C. By contrast, cells incubated in TC regardless of temperature displayed significant and equivalent increases in staining. These data indicate that increased temperature is not the major driver of the mar1Δ cell wall changes.
(A) The mar1Δ cell wall changes are not dependent on temperature. The mar1Δ strain was incubated for 16–18 hours with shaking at the indicated temperature in rich medium (YPD) or tissue culture medium (TC). Exposed chitooligomers in the cell wall were stained with FITC-conjugated WGA and imaged by fluorescent microscopy using the GFP filter. (B) Average fluorescence of at least 100 individual cells was measured using ImageJ/Fiji software. Ns, not significant as determined by one-way ANOVA with Tukey’s multiple comparisons test; all other comparisons, p < 0.0001. (C) The mar1Δ cell wall changes occur with increased pH and can be partially suppressed by glucose supplementation. mar1Δ cells were incubated for 16–18 hours with shaking at 30°C in YPD, YPD buffered to pH 7.4, TC, or TC supplemented with 2% glucose. Cells were stained with FITC-conjugated WGA and imaged as above. Bar, 10 μM. (D) Average fluorescence was measured as above. ****, p < 0.0001; ns, not significant, as determined by one-way ANOVA with Tukey’s multiple comparisons test.
We next tested the role for pH in inducing cell wall changes. Standard YPD medium has a pH of 5–6, while TC medium is buffered to pH 7.4. To test if increased pH could induce the mar1Δ mutant cell wall changes, we buffered YPD to pH 7.4 and measured staining by WGA (Fig 3C and 3D, S1A Fig). We observed a small, but significant increase in chitooligomer staining of mar1Δ cells, suggesting that increased pH is involved in TC-induced cell wall changes. Interestingly, when we buffered YPD to pH 8, we observed increased WGA staining of both WT and mar1Δ cells (S1B Fig).
While higher pH induced increased WGA staining, it did not fully recapitulate the level of staining observed in TC medium. We next tried to suppress the mar1Δ cell wall changes by supplementing TC medium with common nutrients found in rich medium. We observed that when we incubated mar1Δ cells in TC medium supplemented with 2% glucose, the intensity of WGA staining was partially suppressed (Fig 3C and 3D). We did not observe any changes in WT staining or any suppression in TC medium supplemented with 1x complete amino acids (S1A and S1B Fig). Together these data indicate that a combination of increased pH and decreased glucose availability in TC medium induces increased chitooligomer exposure of the mar1Δ mutant.
The mar1Δ cell wall has decreased levels of α- and β-glucan, but not chitin or chitosan
We used several independent methods to further quantify the relative levels of cell wall carbohydrates in our strains. First, we used an enzymatic method  to quantify total chitin and chitosan levels in the mar1Δ and WT cell walls after incubation in TC medium, the condition in which we observe differences in staining intensity. Compared to the WT, the mar1Δ mutant displayed no increase in total chitin or chitosan using this biochemical assay (Fig 2F).
We also quantified our initial staining observations using flow cytometry, demonstrating a striking increase in the mean fluorescence intensity (MFI) of WGA-stained mar1Δ cells compared to WT (Fig 4A). Similar to what we saw by staining, we observed a modest increase in the MFI of CFW-stained mar1Δ cells (Fig 4A). Interestingly, we observed two peaks of EY-stained mar1Δ cells, one corresponding to the mean fluorescence intensity (MFI) of WT stained cells, and a second shifted to a higher MFI (Fig 4A). This suggests a non-homogeneous pattern of chitosan exposure after incubation in host-like conditions in mar1Δ cells. We were unable to accurately assess the relative levels of other cell wall carbohydrates such as α-glucan, β-glucan, and mannoproteins using staining and microscopy methods adapted from other fungal species. In all cases, the staining signal of WT cells was similar to unstained controls (Fig 4A). Therefore, we utilized biochemical methods to quantify cell wall carbohydrate composition. We extracted total cell wall material from our strains and used high-performance liquid chromatography (HPLC) to quantify the levels of glucosamine (chitin and chitosan together), glucose (glucans), and mannose (mannosylated proteins) in these extracts [16,33]. Similar to our enzymatic measurements of chitin and chitosan, we observed no significant difference in the level of glucosamine between the WT and mar1Δ strains (Fig 4B). However, by HPLC, we measured a significant decrease in the levels of glucose and mannose in the mar1Δ mutant compared to the WT (Fig 4B). Together, these results suggest that Mar1 is required to maintain normal levels of the outer glucan and mannan cell wall layers. Accordingly, in the absence of functional Mar1, the resulting changes in outer cell wall carbohydrate abundance result in increased exposure, but not total levels, of the inner cell wall chitooligomers, chitin and chitosan.
(A) The mar1Δ cell wall has increased chitin and chitosan staining by flow cytometry, but mar1Δ staining for α-glucans, β-(1,3)-glucan, and mannoproteins is limited above baseline. WT and mar1Δ cultures were incubated for 16–18 hours at 37°C in TC medium, fixed, labeled, and analyzed by flow cytometry. WGA was used to stain exposed chitin and CFW was used to stain total chitin. EY was used to stain chitosan. An MOPC-104E antibody with an anti-mouse AlexaFluor 488 secondary antibody was used to label α-glucan. An Fc-Dectin-1 fusion protein coupled with an anti-human AlexaFluor 488 secondary antibody was used to label β-(1,3)-glucan. Concanavalin A conjugated to AlexaFluor 488 was used to label mannoproteins. Relevant events were gated in the FSC/SSC plots and are represented as histograms with mean fluorescence on the x-axis and cell counts on the y-axis. Unstained cells were sorted as controls to determined positive labeling. (B) The mar1Δ cell wall has decreased glucan and mannan. Cells were incubated for 16–18 hours at 37°C in TC medium, followed by cell wall isolation. Cell wall carbohydrate levels were quantified using high performance anion-exchange chromatography with pulse ampherometric detection (HPAEC-PAD). All data represent means of results from 3 independent cell wall preparations for each strain. *, p < 0.05; **, p < 0.01 as determined by one-way ANOVA with Tukey’s multiple comparisons test. All other comparisons, not significant (C) Cell wall genes are differentially regulated in mar1Δ. A concentration of 107 cells/ml in 25 ml YPD (30°C) or TC (37°C) were incubated for 1.5 hours, followed by RNA extraction and cDNA synthesis. Expression of cell wall biosynthesis genes was determined by real-time PCR. Fold change was calculated relative to WT YPD levels and normalized to the expression of an internal control. Data represent means of results from 2 independent C. neoformans cultures and RNA extractions per strain. *, p < 0.05; **, p < 0.01; ***, p < 0.001; ****, p < 0.0001 as determined by two-way ANOVA with Tukey’s multiple comparisons test.
Mar1 regulates α- and β-glucan synthase expression
To determine the site of regulation for these cell wall changes, we performed quantitative real time PCR analysis of selected cell wall synthesis genes in the WT and mar1Δ strains. Interestingly, as the WT strain transitions from a rich medium to tissue culture medium, there is a marked increase in the expression of genes encoding α- and β-glucans (Fig 4C). In contrast, there is no significant transcriptional change in most chitin synthase and chitin deacetylase genes during this environmental transition, except for the CHS4 chitin synthase. Consistent with our chitooligomer cell wall staining and quantification data, the expression of these chitin synthase and chitin deacetylase genes was not significantly different between WT and mar1Δ in tissue culture medium. By contrast, the genes encoding the major glucan synthases, including the α-glucan synthase AGS1 and the β-1,3-glucan synthase FKS1, displayed statistically significantly decreased expression in tissue culture medium in mar1Δ compared to WT. These data indicate that Mar1 is required for the transcriptional induction of glucan cell wall genes, including AGS1 and FKS1, that are typically upregulated during host-like conditions.
Rim and cell wall integrity pathway signaling is intact in the mar1Δ background
We chose to more fully explore the relationship of Mar1 function with the Rim and CWI signaling pathways given our observation that the mutant strains associated with each of these pathways most closely shared cell wall-related phenotypes with the mar1Δ strain. C. neoformans strains defective in Rim pathway signaling are unable to grow well at alkaline pH or in the presence of elevated salt concentrations, similar to the mar1Δ strain. This signaling pathway is responsible for activating the Rim101 transcription factor, which in turn regulates the expression of many genes required for cell wall integrity and organization during growth in host-like conditions. Additionally, Rim pathway mutants have increased chitooligomer exposure .
The MAR1 gene was not identified as a Rim101 target in previously published comparative transcriptional profiling experiments . However, direct analysis of the raw data from these experiments (NCBI GEO database accession number GSE43189) indicated that the MAR1 locus was not included in the statistical analyses due to its relatively low transcript abundance. To more definitively determine whether MAR1 is a downstream target of the Rim101 transcription factor, we first performed quantitative real time PCR to measure the expression of MAR1 in the WT background in YPD and TC media. We observed that MAR1 had significantly induced expression in TC medium compared to YPD (p = 0.0017) (S2A Fig). To next determine if the expression of MAR1 is regulated in a Rim101-dependent manner, we analyzed updated transcriptional profiling of the rim101Δ strain in TC medium recently carried out in our laboratory (NCBI GEO database accession number GSE110723). Expression of the MAR1 transcript in the rim101Δ mutant versus WT was only modestly reduced (log2-fold change = 0.772, p = 0.04), suggesting that MAR1 is likely not a major target of Rim101.
We also assessed whether Mar1 might function in the Rim signaling pathway upstream of Rim101. Rim pathway activation occurs in a pH-dependent manner, culminating in the cleavage of the Rim101 transcription factor and its translocation to the nucleus . Rim101 cleavage and nuclear localization are disrupted in mutants of upstream Rim pathway activators . In contrast, Rim101 cleavage/activation is intact in the mar1Δ background (S2B Fig). Together these data indicate that Mar1 likely operates in cellular processes independent of the C. neoformans Rim pathway, despite phenotypic similarities of these mutant strains.
The PKC/CWI pathway is responsible for the activation of the Mpk1 MAP kinase protein, which in turn coordinates enhanced cell wall stress resistance. Accordingly, mutants in this pathway display defective phosphorylation of Mpk1. We therefore assessed Mpk1 phosphorylation in response to cell wall stress in the WT and mar1Δ mutant strains. Western blots of total cell lysates from these strains were analyzed using an antibody directed against phosphorylated (activated) Mpk1 [36,37]. After 3.5 hours of incubation in YPD, WT and mar1Δ cells had a modest level of phosphorylated Mpk1 (S3A Fig). After incubation in TC medium, both WT and mar1Δ displayed enhanced Mpk1 phosphorylation (S3A Fig). These results demonstrate that Mar1 is not required for CWI integrity pathway activation under host-like tissue culture conditions as measured by Mpk1 phosphorylation. To further test this, we constructed a mar1Δ mpk1Δ double mutant and analyzed chitooligomer exposure by WGA staining. In agreement with the mar1Δ mutation inducing distinct cell wall changes, the mar1Δ mpk1Δ double mutant displayed more WGA staining then either mpk1Δ or mar1Δ single mutants (S3B Fig). The patterns of Rim and CWI signaling pathway activation in the mar1Δ strain suggest that the Mar1 protein regulates cell wall integrity in a novel manner.
Mar1 localizes to intracellular and cell surface puncta
To better understand the function of Mar1, we generated an N-terminally tagged GFP-Mar1 fusion protein and analyzed its localization by microscopy. We expressed this fusion protein under the constitutively active HIS3 promoter in the mar1Δ mutant strain background. This fusion protein was functional, displaying partial suppression of mar1Δ dry colony morphology on pH 8 plates. After overnight incubation in YPD medium, GFP-Mar1 localized throughout the cell to small punctate structures on endomembranes and at the cell surface (Fig 5A). 3D-projection of Z-stacked images indicated that many of the observed puncta were located on the cell surface (Fig 5B). Compared to what was observed in YPD-incubated cells, the number of GFP-Mar1 puncta was decreased after incubation in TC medium (Fig 5A and 5B). These puncta also appeared larger and more globular in nature, and overall intracellular endomembrane staining was less intense after incubation in TC medium.
(A) Localization of a Mar1-GFP fusion protein was assessed after cells were incubated for 16–18 hours in YPD (30°C) or TC (37°C). Live cells were imaged using DeltaVision deconvolution fluorescent microscopy with the GFP filter. Images were deconvolved using softWoRx software. Bar, 10 μM. (B) Cells were incubated and imaged as above. 3D-projections were generated from z-stacked images using ImageJ/Fiji software and are pseudo-colored to indicate brightness of staining from yellow to blue. Bar, 10 μM. (C) A Mar1-GFP expression strain was pre-incubated in YPD medium to mid-log phase and transferred to TC medium at time zero. Cells were incubated for the indicated time in TC at 37°C with shaking and imaged by fluorescent microscopy using the GFP filter. Bar, 10 μM. (D) Quantification of staining by FITC-conjugated WGA. The WT and mar1Δ strains were incubated to mid-log phase in YPD medium and transferred to TC medium at time zero. Exposed chitooligomers in the cell wall were stained with FITC-conjugated WGA, imaged by fluorescent microscopy using the GFP filter, and average fluorescence was quantified by ImageJ/Fiji software. ****, p < 0.0001 as determined by two-way ANOVA with Sidak’s multiple comparisons test.
To further characterize the dynamics of GFP-Mar1 protein localization, we analyzed protein fluorescence over time after shifting cells to TC medium (Fig 5C). After 2 and 4 hours in TC medium, GFP-Mar1 localizes to small punctate structures as well as to endomembranes, similar to what we observed for cells incubated in YPD overnight. After 6 hours, some GFP-Mar1 puncta begin to appear globular, more similar to what we observed for cells incubated in TC overnight. By 8 hours, the majority of GFP-Mar1 puncta appear globular. Interestingly, when we measured mar1Δ chitooligomer exposure over time in TC medium, we observed a similar time course of changes (Fig 5D). Increased WGA staining of mar1Δ cells can be observed beginning around 6 hours after incubation in TC medium, similar to when globular GFP-Mar1 puncta are seen.
Intracellular trafficking is impaired in mar1Δ
The dynamic localization of GFP-Mar1 led us to next investigate intracellular trafficking processes in the mar1Δ mutant strain. To determine if these processes were impaired, we used the lipophilic dye FM4-64 to assess rates of endocytosis in YPD and TC media (Fig 6A). Medium-sized, bright endocytic vesicles can be seen after 30 minutes of staining in WT and mar1Δ cells incubated in YPD. Endocytic vesicles can also be observed in WT cells incubated in TC medium; However, by contrast, less well-defined, tubular structures are observed in mar1Δ cells incubated in TC medium.
(A) Uptake of the lipophilic dye, FM4-64, is irregular in mar1Δ cells. Cells were incubated overnight in YPD (30°C) or TC (37°) and stained with FM4-64 for 30 minutes with shaking, followed by pelleting and refreshing in the indicated media for an additional 30 minutes with shaking. Stained cells were imaged by fluorescent microscopy with the Texas Red filter and were taken with the same exposure time. Bar, 10 μM. Arrows indicate endocytic vesicles (B, C) Acid phosphatase secretion is intact in the mar1Δ strain; alkaline phosphatase activity is decreased. WT and mar1Δ cultures were pre-incubated for 16–18 hours in phosphate replete minimal medium. The cells were diluted to an OD of 0.9 in either phosphate replete (non-inducing) or phosphate deficient (inducing) minimal medium and incubated for 3 hours at 30°C with shaking. (B) Acidic or (C) alkaline para-Nitrophenylphosphate (pNPP) substrate solution was added to each well and plates were incubated for an additional 2.5 hours at 37°C with shaking. Phosphatase activity was measured as absorbance at 410 nm over cell density at 600 nm. Data represent the mean of 3 replicates per strain per condition. (D) Fks1 is mislocalized in TC medium in mar1Δ cells. The Fks1-GFP fusion protein was expressed in the WT and mar1Δ mutant strains. Cells were incubated for 16–18 hours in TC medium at 37°C. Live cells were imaged using DeltaVision deconvolution fluorescent microscopy with the GFP filter. Images were deconvolved using softWoRx software. Bar, 10 μM.
To further assess intracellular trafficking, we measured acid and alkaline phosphatase activity in cell supernatants. In S. cerevisiae, acid phosphatase is secreted through the canonical secretory pathway, whereas alkaline phosphatase is trafficked to the vacuolar membrane through the alternative ALP pathway . Both enzymes are induced under low phosphate conditions and can be assayed by measuring the colorimetric hydrolysis of the para-Nitrophenylphosphate (pNPP) substrate. Over a time-course of incubation with the pNPP substrate in acidic medium, phosphate starved (induced) WT and mar1Δ cells showed increased phosphatase activity compared to phosphate replete (non-induced) cells (Fig 6B). A similar induced increase in phosphatase activity was observed for WT cells in alkaline conditions; however, there was a minimal difference in phosphatase activity between phosphate starved and phosphate replete mar1Δ cells in alkaline conditions, suggesting a defect in secreted alkaline phosphatase activity (Fig 6C). Together, these data are consistent with defects in endocytic and vesicular trafficking in the mar1Δ mutant, but not a defect in classical secretion.
Localization of the Fks1 β-(1,3)-glucan synthase is altered in mar1Δ
To further explore the mar1Δ mutant defect in glucan abundance, we constructed a C-terminally tagged Fks1-GFP fusion protein. We transformed this allele into the WT H99 background and selected those transformants in which the allele replaced the endogenous locus for further analysis. Previous work has indicated that FKS1 is an essential gene in C. neoformans . Therefore, the ability to replace the WT FKS1 allele with the FKS1-GFP allele suggests that this fluorescent fusion protein is functional. We subsequently mutated the MAR1 gene in the strain expressing Fks1-GFP, and we analyzed the localization of Fks1-GFP in two independent mar1Δ mutants. After incubation in YPD medium, we observed by fluorescent microscopy that Fks1-GFP localizes to punctate structures on cellular membranes in both the WT and mar1Δ backgrounds (S4A Fig). After incubation in TC medium, Fks1-GFP localization is more heterogeneous among the cell population. It is still present in puncta throughout the cell in the majority of cells, but it is enriched at the plasma membrane of some cells in the WT background, predominately mother cells (Fig 6D). By contrast, Fks1-GFP localization is more homogeneous among cells in the mar1Δ background, with reduced plasma membrane puncta and very few cells displaying the uniform plasma membrane localization observed in the WT (Fig 6D and S4B Fig). These data suggest that Mar1 is required for proper trafficking and localization of Fks1 to the cell membrane in TC medium.
Cell wall ultrastructure and vesicular trafficking appear disordered in mar1Δ cells
To better visualize the cell wall changes of the mar1Δ mutant, we examined our strains by transmission electron microscopy (TEM) after incubation in either YPD or TC media. In rich YPD medium, both WT and mar1Δ cells had thin, organized cell walls with a well-defined lamellar appearance (Fig 7A). The mar1Δ cell walls displayed a trend to be slightly thinner than WT cell walls in this condition, but this difference was not statistically significant (p = 0.3431). In TC medium, the cell walls of WT cells remained compact and well-ordered (Fig 7B). In contrast, mar1Δ cell walls were less compact and well-organized after incubation in TC medium and significantly thicker than WT or mar1Δ cell walls incubated in YPD (p = 0.0011 and 0.0007 respectively). Several cells appeared to have layers of cell wall material sloughing away from the cell periphery, a phenotype that was not observed for WT cells (Fig 7B). Accordingly, a significant amount of debris was also observed in the space between cells in the mar1Δ TC samples, likely representing degraded cell wall material, or perhaps changes in chemical cross-linking resulting in altered cell wall integrity during sample processing.
WT and mar1Δ cells were incubated for 16–18 hours in YPD (30°C) and TC (37°C) medium, followed by glutaraldehyde and KMnO4 fixation and partial dehydration as described previously [14,68,101]. Samples were further processed, embedded, sliced, and imaged by transmission electron microscopy (TEM). (A) YPD incubated cells display thin, ordered cell walls. Left image of each pair: bar, 1 μM. Right image of each pair: bar, 200 nm. Letters indicate location of inset. (B) mar1Δ cells incubated in TC medium have a less organized cell surface. Left image of each pair: bar, 1 μM. Right image of each pair: bar, 200 nm. Letters indicate location of inset. Red arrows indicate apparent cell wall material disassociating from cell surface. (C) There are increased numbers of electron lucent structures in mar1Δ cells incubated in TC medium. Bar, 1 μM. Red asterisks indicate mar1Δ cells with several electron lucent vesicles near cell periphery.
In addition to the notable TC-induced cell wall changes in the mar1Δ strain, the TEM images also suggested alterations in vesicular trafficking. In both WT and mar1Δ strains, there were numerous membrane-bound vesicles. These structures have been suggested to be secretory vesicles transporting cargo toward the cell surface . In TC medium, both the WT and mar1Δ strains demonstrated a relative increase in the number of these vesicles carrying electron-dense material localized near the cell surface. In mar1Δ cells, an increase in large electron-lucent vesicles was also observed (Fig 7C). In some cells, multiple large “empty” vesicles can be seen near the cell periphery. In S. cerevisiae, the accumulation of similar vesicles has been observed for several secretory mutants [41,42]. Similarly, the sav1Δ secretory mutant in C. neoformans accumulates post-Golgi secondary vesicles . Along with the altered FM4-64 staining and alkaline phosphatase secretion described, these images support a defect in intracellular trafficking in this mutant.
The mar1Δ mutant cell wall changes are associated with increased recognition by macrophages and dendritic cells
Macrophages and dendritic cells are likely the first immune cells that C. neoformans contacts within infected lungs. To determine if the changes in the mar1Δ cell wall would affect this interaction, we quantified the production of tumor necrosis factor-alpha (TNF-α) after co-culturing WT and mar1Δ strains with primary bone marrow-derived macrophages (BMMs) and bone marrow-derived dendritic cells (BMDCs). We have previously used this pro-inflammatory cytokine as a marker of macrophage activation in vitro, observing that C. neoformans strains with increased chitooligomer exposure often induce more TNF-α production than WT [14,16]. Accordingly, we observed that mar1Δ cells induced significantly more TNF-α production from both BMMs and BMDCs, compared to the WT or reconstituted strains (Fig 8A and 8B). To determine if this is due to an active cellular process, we tested the macrophage response after co-culturing with heat-killed mar1Δ cells. Similar to live cells, we found that heat-killed mar1Δ cells also induce increased TNF-α production by BMMs, demonstrating that mar1Δ macrophage stimulation is not dependent on cell viability (Fig 8A).
(A) Macrophage activation by mar1Δ cells is not dependent on cell viability. WT, mar1Δ, and mar1Δ + MAR1 cells were incubated for 16–18 hours at 37°C in TC medium. Bone marrow-derived macrophages (BMMs) were co-incubated with live or heat-killed (HK, 1–2 hours at 65°C) C. neoformans strains at a multiplicity of infection (MOI) of 10:1, C. neoformans:BMMs. TNF-α levels (pg/ml) were assayed from the supernatant by ELISA. Data represent means from 3 replicates per strain per condition from 3 independent experiments (n = 9). *, p < 0.05; ***, p < 0.001 mar1Δ vs. WT as determined by two-way ANOVA with Tukey’s multiple comparisons test. (B) mar1Δ induces increased TNF-α production by dendritic cells. C. neoformans cells were incubated as described above and co-incubated with bone marrow derived dendritic cells (BMDCs) at an MOI of 10:1, C. neoformans:BMDCs. TNF-α levels were assayed from the supernatant by ELISA. Data represent means from 3 replicates per strain from 2 independent experiments (n = 6). **, p < 0.01 mar1Δ vs. WT as determined by one-way ANOVA with Tukey’s multiple comparisons test. (C) Macrophage activation by mar1Δ cells is dependent on pre-culturing in TC medium. Cells were incubated for 16–18 hours in YPD (30°C) or TC (37°C) prior to co-incubation with BMMs and TNF-α quantification by ELISA as above. Data represent means from 3 replicates per strain (n = 3). ****, p < 0.0001 mar1Δ/TC vs. WT/ TC as determined by two-way ANOVA with Tukey’s multiple comparisons test. mar1Δ/YPD vs. WT/YPD, not significant. (D) Cell wall material isolated from mar1Δ induces increased macrophage activation. WT, mar1Δ, and mar1Δ + MAR1 reconstituted cells were incubated for 16–18 hours at 37°C in TC medium, followed by cell wall isolation. 10 μg/ml of cell wall material was co-cultured with BMMs and TNF-α was quantified from the supernatant by ELISA. Data represent means of 3 independent cell wall isolations in 3 independent experiments (n = 9 for each strain). *, p < 0.05 mar1Δ vs. WT as determined by one-way ANOVA with Tukey’s multiple comparisons test.
Due to our observation that the mar1Δ cell wall changes are only induced when the cells are incubated in host-mimicking tissue culture conditions, we tested both YPD- and tissue culture media-incubated cells in our BMM assays. Consistent with the host-induced changes in the mar1Δ cell wall, only cells pre-incubated in TC medium prior to co-culture induced increased TNF-α production from BMMs (Fig 8C). This was the case for both live and heat-killed cells. We also examined whether the cell wall itself was sufficient to elicit a response from BMMs. Compared to WT and reconstituted strain controls, we observed that isolated cell walls from the mar1Δ strain pre-incubated in TC medium induced more TNF-α production by BMMs (Fig 8D). Together these data suggest that macrophage activation by this strain is dependent on specific cell wall changes that are induced by host-mimicking tissue culture conditions.
Cell wall changes in mar1Δ increase recognition of the acapsular cap59Δ mutant
The polysaccharide capsule can serve to shield the immunogenic cell surface of C. neoformans from immune recognition. Additionally, capsule polysaccharide itself can be immunosuppressive . We have observed that the mar1Δ mutant does not properly attach capsule to its cell surface, but it seems to secrete the polysaccharide similarly to WT (Fig 2A and 2B). Therefore, we wanted to determine if the capsule-deficient phenotype of mar1Δ is contributing to its ability to stimulate macrophages. To test this, we generated a mar1Δ cap59Δ double mutant; Cap59 is involved in capsule biosynthesis, and the cap59Δ mutant does not produce any detectable capsule polysaccharide [14,44]. We used the cap59Δ single mutant as our control strain, and tested macrophage activation of the mar1Δ cap59Δ double mutant compared to cap59Δ alone. We observed that the mar1Δ cap59Δ double mutant induced significantly more TNF-α than the single cap59Δ mutant (S5 Fig). This indicates that the cell surface of mar1Δ has a role in activating macrophages that is independent of this strain’s capsule defect and separate from any cell surface changes that the cap59Δ mutant exhibits as a result of its inability to produce capsular polysaccharide.
Mar1 is required for full virulence in vivo
We used a murine inhalation model of cryptococcosis to assess the role of Mar1 in virulence . We intranasally inoculated C57BL/6 mice with WT, mar1Δ, or mar1Δ + MAR1 complemented cells and monitored mice over the course of 40 days for clinical endpoints predictive of mortality. Mice infected with the WT or mar1Δ + MAR1 complemented strains exhibited a median survival time of 18 days (Fig 9A). In contrast, mice infected with the mar1Δ mutant had a median survival time of 28 days. We also measured fungal burden in the lungs at early time points after infection. As early as days 1 and 4 after infection, the number of mar1Δ mutant cells were significantly decreased in the lungs of infected mice compared to WT cells (Fig 9B). However, half of the mar1Δ-infected mice eventually succumbed to infection, suggesting there is not complete clearance of these cells.
(A) The mar1Δ strain is attenuated in the C57BL/6 mouse background. For each strain, 9–10 C57BL/6 mice were inoculated with 1 x 105 cells, monitored daily for signs of infection, and sacrificed at predetermined clinical endpoints that predict mortality. Statistical significance was determined by log-rank test with Bonferroni correction: p < 0.0001, mar1Δ vs. WT; p < 0.001, mar1Δ vs. mar1Δ + MAR1; not significant. (B) There are minimal mar1Δ cells in the lungs of infected mice at early time points. Colony forming units (CFUs) were determined from lung homogenates from 5 mice per strain at days 1 and 4 post inoculation. *, p < 0.05 mar1Δ vs. WT; **, p < 0.01 mar1Δ vs. mar1Δ + MAR1 as determined by one-way ANOVA with Tukey’s multiple comparisons test. (C) The mar1Δ strain is avirulent in the BALB/c mouse background. For each strain, 9–10 BALB/c mice were inoculated with 1 x 105 cells, monitored daily, and sacrificed as described above. Statistical significance was determined by log-rank test with Bonferroni correction: p < 0.0001, mar1Δ vs. WT/mar1Δ + MAR1; not significant, WT vs. mar1Δ + MAR1. (D) The mar1Δ strain is not completely cleared from BALB/c mice despite mouse survival. CFUs were determined from lung and brain homogenates from mar1Δ infected mice post-mortem. Ratio indicates number of mice represented.
It has been previously documented that mice from different genetic backgrounds display varying levels of sensitivity to cryptococcal infection [46,47]. C57BL/6 mice predominate towards protective Th1 type responses, while BALB/c mice have a non-protective Th2 type bias . Therefore, we also tested the susceptibility of BALB/c mice to mar1Δ infection. We intranasally inoculated BALB/c mice as above and monitored mice over the course of a 40-day infection. In contrast to C57BL/6 mice, all of the mar1Δ-infected BALB/c mice survived the course of the experiment (Fig 9C). At the end of the experiment, we measured fungal burden in the lungs and brain post-mortem and observed that mar1Δ is not completely cleared from all mice, despite mouse survival (Fig 9D).
In vitro response to mar1Δ requires members of the C-type lectin receptor and Toll-like receptor families
The initial innate interaction between fungi and the host begins with pathogen recognition by pattern recognition receptors (PRRs) on innate immune cells. Members of the C-type lectin receptor (CLR) and Toll-like receptor (TLR) families have been implicated in recognizing fungal pathogen-associated molecular patterns (PAMPs), including fungal cell wall components. Many of the CLRs signal through the adaptor protein, Card9, to downstream cellular pathways to activate pro-inflammatory cytokines. Similarly, several of the TLRs signal through the adaptor protein, MyD88, to activate downstream cellular responses.
To determine if members of the CLR or TLR families are responsible for recognizing the mar1Δ cell surface, we examined the roles of the Card9 and MyD88 adaptor proteins in macrophage activation by these strains. We co-cultured our strains with BMMs isolated from Card9-/- and MyD88-/- mice and compared the production of TNF-α with BMMs isolated from WT mice. Consistent with our results above, the mar1Δ mutant induced significantly more TNF-α production from WT BMMs than isogenic WT C. neoformans strains. By contrast, the Card9-/- BMMs had a significant, but incomplete, reduction in their response to the mar1Δ mutant, indicating a role for both Card9-dependent and–independent receptors (Fig 10A). Strikingly, the exaggerated TNF-α response to the mar1Δ strain was completely absent in MyD88-/- BMMs, suggesting a significant role for TLRs in recognizing the mar1Δ cell surface (Fig 10B). Combined, these data demonstrate that both CLR and TLR family receptors are likely involved in sensing and responding to the mar1Δ strain.
BMMs were harvested from the indicated mouse strains (C57BL/6 background, unless otherwise noted) and co-incubated with C. neoformans (Cn) cells at an MOI of 10:1, Cn:BMMs, followed by quantification of TNF-α (pg/ml) in the supernatant by ELISA. (A) Card9 is involved in macrophage activation by mar1Δ cells. Data represent means of 3 replicates from 3 independent experiments (n = 9). **, p < 0.01 WT vs. Card9-/- BMMs as determined by two-way ANOVA with Sidak’s multiple comparisons test. (B) MyD88 is required for TNF-α production by macrophages in response to mar1Δ. Data represent means of 3 replicates from 3 independent experiments (n = 9). ****, p < 0.0001 WT vs. MyD88-/- BMMs as determined by two-way ANOVA with Sidak’s multiple comparisons test. (C) Dectin-1 is involved in the response to mar1Δ cells. Data represent means of 3 replicates from 2 experiments (n = 6). ****, p < 0.0001 WT vs. Dectin-1-/- BMMs as determined by two-way ANOVA with Sidak’s multiple comparisons test. (D) TLR2/4-/- BMMs do not respond to mar1Δ cells. Data represent means of 3 replicates (n = 3). ****, p < 0.0001 WT vs. TLR2/4-/- BMMs as determined by two-way ANOVA with Sidak’s multiple comparisons test. (E) TLR4 is not required for the production of TNF-α induced by mar1Δ cells. BMMs were isolated from C3H/HeJ mice with a null mutation in TLR4, and C3H/HeOuJ control mice. Data represent means of 3 replicates from 2 independent experiments (n = 6). **, p < 0.01 C3H/HeOuJ vs. C3H/HeJ BMMs as determined by two-way ANOVA with Sidak’s multiple comparisons test. (F) Macrophage activation by mar1Δ is partially dependent on TLR2. Data represent means of 3 replicates from 3 independent experiments (n = 9). ****, p < 0.0001 WT vs. TLR2-/- BMMs as determined by two-way ANOVA with Sidak’s multiple comparisons test.
Recognition of the mar1Δ cell surface involves multiple PRRs important for chitin sensing
Based on the importance of both Card9 and MyD88 in sensing and responding to mar1Δ cells, we next focused on specific PRRs that have been previously implicated in fungal cell wall recognition, in particular chitin and chitooligomers. The C-type lectin receptor, Dectin-1 has been studied extensively for its role in recognizing the fungal cell surface, in particular β-glucan in the cell wall . It has also been implicated in sensing fungal chitin and leading to a pro-inflammatory response, including the production of TNF-α, from innate immune cells [19,20]. To test the role of Dectin-1 in recognizing mar1Δ cells, we co-cultured our strains with Dectin-1-/- BMMs and measured the production of TNF-α after 6 hours (Fig 10C). Compared to WT BMMs, the response to mar1Δ by Dectin-1-/- BMMs was significantly decreased, indicating that this C-type lectin receptor is involved in sensing mar1Δ.
Due to the marked decrease in the mar1Δ response by MyD88-/- BMMs, we used BMMs isolated from TLR2/4-/- mice to focus on the extracellular TLRs that have been previously implicated in fungal PAMP recognition [50–52]. We co-cultured our strains with TLR2/4-/- BMMs and measured the TNF-α response after 6 hours compared to WT BMMs. Similar to what we observed for MyD88-/- BMMs, TLR2/4-/- BMMs showed significantly reduced responses to mar1Δ, indicating a role for TLR2 and/or TLR4 in sensing this fungal cell surface (Fig 10D).
To test the individual role of TLR4, we utilized C3H/HeJ mice that have a null mutation in the TLR4 gene . These mice have been used in several studies to assess the role of TLR4 in the response to a variety of microbial pathogens or pathogen products including bacteria (Mycobacterium , Bordetella , Rickettsia , Neisseria ) and fungi (Coccidioides , Aspergillus ). We isolated BMMs from C3H/HeJ mice and C3H/HeOuJ control mice and assessed their TNF-α responses to mar1Δ. We found a modest increase in the response to mar1Δ between C3H/HeJ TLR4 mutant BMMs and C3H/HeOuJ control mice (Fig 10E). In control experiments, both mutant and control BMMs responded similarly to a control ligand, zymosan; however, as expected, C3H/HeJ BMMs did not respond to the canonical TLR4 ligand, LPS, while C3H/HeOuJ BMMs responded normally (S6 Fig). These results suggest that the response to mar1Δ is TLR4-independent.
In previous work, TLR2 has been described to have a role in recognizing fungal chitin, leading to a pro-inflammatory response and the production of TNF-α [18–20]. To test if TLR2 is responsible for recognizing the mar1Δ surface, we carried out our co-culture assays using TLR2-/- BMMs. The TNF-α response after 6 hours was significantly reduced in the TLR2-/- BMMs compared to WT BMMs (Fig 10F). These data suggest that the decrease in response to mar1Δ in the TLR2/4-/- BMMs was due to the defect in TLR2. Together these results support a model in which the TLR2 and Dectin-1 PRRs are both involved in sensing and responding to the mar1Δ cell surface.
The fungal cell wall is a dynamic structure that is constantly being remodeled. Fungal pathogens carefully regulate their cell wall in the context of the host in order to adapt to this environment, as well as to avoid detection by the immune system. Here we have identified a novel cell wall remodeling protein in C. neoformans, Mar1, and demonstrated that this protein has a role in intracellular trafficking that results in the proper localization of the cell wall β-(1,3)-glucan synthase, Fks1, as well as the continuous reorganization of the fungal cell surface in host-like conditions. The immunological consequence of this cell wall mis-regulation in the mar1Δ mutant is an increased activation of macrophages and dendritic cells as well as attenuated virulence in vivo.
Our lab and others have demonstrated that the cell wall changes in size and in composition in response to the host environment [14–16,60]. Here, in WT cells, we have observed an increase in cell wall thickness in TC medium by TEM. This occurs in the setting of the well characterized formation and expansion of polysaccharide capsule. We have also measured increased expression of many of the known cell wall synthase genes in response to TC medium, and we have demonstrated that Fks1 is enriched at the plasma membrane in a specific population of WT cells after incubation in TC medium. By contrast, we observed a thickened, but less structurally sound cell wall in mar1Δ cells incubated in TC medium. Despite this observed increase in cell wall thickness, we showed that the mar1Δ strain has decreased total levels of glucans and mannans, decreased expression of glucan synthase genes, and decreased Fks1 protein at the plasma membrane in TC-incubated cells. We also demonstrate that mar1Δ cells do not properly attach capsule to their cell surface. Together these data suggest a model in which the mar1Δ strain has decreased total levels of outer cell wall components (glucans and mannans), leading to the observed capsule attachment defect and increased exposure of the inner cell wall components chitin and chitosan (Fig 11).
When incubated in tissue culture medium, there is an increase in the expression of genes encoding the biosynthesis enzymes for outer cell wall components, resulting in a thicker cell wall, enhanced capsule attachment, and immune avoidance. The Mar1 protein is required for controlling aspects of these cell wall adaptations, including the proper induction and localization of glucan synthases. As a result, in the mar1Δ mutant the levels of these outer cell wall components are reduced, exposing the more immunogenic chitooligomers that are normally masked from immune recognition.
Mar1 is impacting cell wall remodeling through regulation of the cell wall synthase response to host-like conditions
Our model demonstrates a prominent role for Mar1 in the cell wall response to TC medium, in particular through the regulation of cell wall synthase expression and localization. Previous studies in model fungi have shown differential localization of cell wall synthases in response to different conditions. The S. cerevisiae chitin synthase, Chs3, is maintained under normal conditions in internal compartments called “chitosomes”, where it is cycled between early endosomes and the trans-Golgi network in a clathrin adaptor protein (AP) complex dependent manner . However, under stress conditions, Chs3 is trafficked to the plasma membrane in order to synthesize increased chitin . Similar clathrin AP complex-dependent trafficking has been described for the Schizosaccharomyces pombe β-(1,3)-glucan synthase, Bgs1. Mutations in the conserved AP-1 adaptor complex protein, Sip1, and the clathrin light chain protein, Clc1, lead to defects in Bgs1 localization/delivery to the plasma membrane [63,64].
Here we have observed that Fks1 plasma membrane localization is enriched in specific WT cell populations upon shifting cells to tissue culture medium, suggesting a similar stress-induced cycling of this protein to its functional location at the cell surface. Mar1 could be impacting the cycling of Fks1 between internal compartments and the plasma membrane in multiple ways. For example, Mar1 may be required for exocytic movement towards the plasma membrane, and therefore in the absence of this protein, exocytosis and/or secretion of Fks1 is defective. However, the fact that capsule, melanin, and acid phosphatase secretion remain intact in this strain would suggest that Mar1 is not impacting generalized exocytosis or secretion. Another way in which Mar1 could be impacting Fks1 trafficking is if it were required for protein maintenance on the PM, for example, by preventing excessive endocytosis or recycling. Our data showing altered endocytosis of FM4-64 in the mar1Δ strain would support this model.
Another alternative mechanism by which Mar1 might impact Fks1 localization and ultimately cell wall remodeling could be through a role in cell surface signaling. The Mar1 protein has two transmembrane domains and our microscopy data indicates that it localizes to cellular membranes at the cell surface. Therefore, it is feasible to hypothesize that Mar1 is serving a signaling role, perhaps as a cell surface receptor, propagating the cue to traffic cell wall synthase proteins from internal stores to the PM under stress conditions. This model for Mar1 function, unlike the previous, would also explain the lack of transcriptional response of the FKS1 and AGS1 genes in the mar1Δ background. If the mutant cells are unable to sense specific stress conditions, it is plausible that they would not induce transcription of the enzymes required to respond to that stress.
The differential localization of Mar1 in YPD and TC media is also consistent with Mar1 serving a sensor/receptor role. We observed many Mar1 puncta, particularly on the cell surface, but also on intracellular membranes resembling the endoplasmic reticulum when the cells were incubated in YPD medium. By contrast, when shifted to TC medium, we observed a decrease in the number of puncta, as well as changes in their size/shape. It is possible that this decrease in puncta represents the endocytosis of Mar1 from the membrane to transmit a signal downstream, or perhaps internalization for degradation. Interestingly, we observed these changes in Mar1-localization at similar time points as increases in cell wall chitooligomer staining in the mar1Δ mutant; these observations suggest a correlation between Mar1 localization and the resulting cell wall response.
A role for Mar1 as a sensor implies a particularly intriguing model, as few cell surface stress sensors have been identified by sequence homology in C. neoformans. Interestingly, the putative Rra1 pH sensor was recently identified in C. neoformans; while appearing structurally and functionally similar to that of the pH sensors in ascomycete fungi, it was found to share no significant sequence similarity . While Mar1 does not share sequence similarity with any known sensor proteins, its structural features and pattern of dynamic localization suggest that it could be serving as a functional homologue of a cell stress sensor.
Proper cell wall regulation is required for effective immune avoidance
The importance of cell wall remodeling, in particular as it relates to cell wall masking, has been well-described in other fungi. The α-glucan layers of H. capsulatum and A. fumigatus have both been implicated in shielding β-glucan in these fungi. β-glucanases that degrade exposed β-glucan have also been elucidated in H. capsulatum . In C. albicans, several factors have been implicated in β-glucan exposure including morphotype switching, antifungal exposure, phospholipid production, and carbon source [5,6,66]. For example, Ballou, et al. recently demonstrated that lactate exposure elicits increased β-glucan masking in C. albicans and other pathogenic Candida species, leading to reduced recognition by immune cells . Furthermore, cell wall remodeling has been shown to drive immune function even after ingestion by host phagocytes. O’Meara, et al. showed that C. albicans actively remodels its cell surface upon phagocytosis in order to induce inflammatory cell death pathways in macrophages to aide in fungal cell escape .
C. neoformans is particularly adept at hiding from the immune system. Importantly, the polysaccharide capsule serves to effectively shield potentially immunostimulatory molecules from host detection [29,67]. However, there are several examples of the immune consequences of improper cell wall organization in C. neoformans. The loss of α-(1,3)-glucan results in a highly disorganized cell wall, with increased chitin and chitosan content and redistributed β-glucan, rendering cells avirulent in mouse models of infection . Chitosan-deficient strains of C. neoformans are also unable to cause disease in mice, inducing a protective proinflammatory host immune response that leads to their rapid clearance [69–71]. Imprecise regulation of the cell wall has also been implicated in excessive immune responses related to improper C. neoformans cell wall exposure, in particular chitooligomers (chitin and/or chitosan). Our laboratory has previously studied the importance of the Rim101 alkaline pH transcription factor in regulating cell wall remodeling [14–16]. The rim101Δ mutant exposes increased cell wall chitooligomers, which results in an excessive and nonprotective immune response in vivo . Likewise, Wiesner, et al. demonstrated that strains with increased chitin abundance induced unfavorable immune outcomes and exacerbated disease .
Here, we propose that both the exposure and total levels of cell wall components are important determinants of immune recognition of fungi. Based on our HPLC data, our model suggests that mar1Δ has less β-glucan in its cell wall (Fig 11). Therefore, one might predict that immune signaling would be less active with reduced immunostimulatory β-(1,3)-glucan to sense. However, the mar1Δ cell wall has more exposed chitooligomers than WT. As noted above, several studies have highlighted the importance of chitin and chitin-derived structures in C. neoformans immune recognition [12,16]. Additionally, cells in which chitooligomer exposure was blocked by WGA exhibited reduced association with murine macrophages . While a single chitin receptor has not been identified to date, several PRRs have been implicated in different aspects of chitin recognition. Here we have demonstrated that multiple PRRs are also involved in the recognition of the mar1Δ cell wall. Macrophage activation by mar1Δ was partially dependent upon Card9, the adaptor protein required for most C-type lectin receptor signaling, and entirely dependent on MyD88, the adaptor protein required for signaling through many Toll-like receptors (TLRs). Accordingly, we demonstrated a requirement for Dectin-1 and TLR2 in the activation of macrophages in response to the mar1Δ mutant strain. There are several possibilities as to why more than one PRR might be involved in this process. In addition to their roles individually, PRRs can function together to induce downstream immune signaling pathways. In particular, several groups have demonstrated a collaborative role between Dectin-1 and TLR2 in detecting fungal epitopes [73–75].
Wagener and colleagues proposed a model in which chitin recognition occurs in different stages during C. albicans interaction with immune cells . During early interaction, chitin on the surface of this fungus is recognized by Dectin-1 and TLR2, resulting in the secretion of proinflammatory TNF-α. Over time this leads to the secretion of chitinases and a decrease in pathogen load. Later in this interaction, digested fragments of chitin are phagocytosed by immune cells and recognized by intracellular TLR9 and NOD2 in a mannose receptor (MR)-dependent manner, resulting in the secretion of IL-10. This latter state occurs during the resolution of infection. In agreement with this model, other groups have demonstrated that intermediate-sized chitin stimulated macrophage production of TNF-α that is dependent on TLR2, Dectin-1, and MR. In contrast, small chitin fragments induced IL-10 production . Given the importance of chitooligomer size and structure in the immunostimulatory process, future work on the mechanism of chitin and chitosan synthesis and degradation will elucidate its role in fungal immune recognition and evasion. Our data demonstrating that intact mar1Δ cells induce increased TNF-α from macrophages in a TLR-2 and Dectin-1 dependent manner support these emerging models of the centrality of chitooligomer exposure in fungal stimulation of the host immune system.
It is still possible, that other factors are contributing to the mar1Δ macrophage activation phenotype. While we demonstrated decreased overall levels of glucans and mannans in mar1Δ cell wall extracts, this does not necessarily exclude these components from playing a role. We did not observe a significant signal from Fc-Dectin-1 labeled cells above baseline, however we did see a very modest increase in mar1Δ cells stained with the α-glucan antibody, MOPC-104E, and the mannoprotein-binding lectin, Concanavalin A (Fig 4A). While α-glucans generally serve to mask other cell wall components, Cryptococcal mannoproteins are well-known to illicit immune responses . We have also shown that mar1Δ cells lack a polysaccharide capsule, and further demonstrated that this is due to a capsule attachment defect, rather than a biosynthesis or secretion defect. Polysaccharide capsule serves to shield the cell surface from immune detection, and free capsule has been shown to have immunomodulatory effects [9,77]. Using a mar1Δ cap59Δ double mutant, in which capsule biosynthesis is inhibited, we showed that the mar1Δ cell wall has an impact on macrophage activation that is independent of the cap59Δ single mutant strain, suggesting this response is not due simply to the lack of capsule in the mar1Δ background. We also determined that heat killed mar1Δ cells, in which capsule polysaccharide is not being actively shed, are still immunostimulatory.
Based on our data, we suggest multiple ways in which this capsule defect may arise. First, it has been previously shown that capsule attachment to the cell wall requires Ags1 expression and α-glucan . We demonstrated that the mar1Δ mutant has decreased glucans in its cell wall and severely decreased Ags1 expression. We also observed that while mar1Δ sheds a similar amount of capsule polysaccharide into the media as WT cells, it migrated at a different rate. This could suggest that the major capsule polysaccharide component, GXM, is modified in some way as to inhibit its ability to attach to the cell wall.
Additional yet unidentified PRRs may also be involved in this interaction. In addition to further interrogating the role of mannoproteins in the mar1Δ response, future work will examine the importance of the mannose receptor in this interaction. The role of TLR9 and NOD2 in this interaction is also intriguing. We did not explore these intracellular receptors in this study, however TLR9 and NOD2 have been previously implicated in recognizing fungal chitin that has been digested and phagocytosed by mammalian macrophages . This recognition leads to the production of the anti-inflammatory cytokine, IL-10, and is thought to lead to the resolution of the pro-inflammatory immune response to intact fungal chitin . In earlier studies, it was observed that, in fact, TLR9-/- macrophages co-incubated with C. albicans and S. cerevisiae had increased TNF-α production, suggesting a role for TLR9 in modulating the pro-inflammatory response to these fungi . TLR9 deficiency has also been associated with worsened immune outcomes in C. neoformans [79,80].
In summary, we have identified a novel cell wall regulatory protein, Mar1. This protein, while not apparent in any canonical cell wall regulatory pathway, is required for proper cell wall organization in host-like conditions. The Mar1 protein localizes to discrete puncta on cellular membranes, with an apparent reduction in puncta upon shifting cells to host-like conditions. In the absence of Mar1, transcription of the α- and β-glucan synthases is not induced, leading to a decrease in these cell wall components and an increase in the exposure of the chitooligomers, chitin and chitosan. We propose that the role of Mar1 in cell wall integrity is in orchestrating the proper response to host-like conditions, occurring in part at the level of intracellular trafficking and results in the mis-localization of the β-(1,3)-glucan synthase, Fks1. Here we have also shown that this dysregulated cell wall manipulates the host-pathogen interaction, leading to increased macrophage activation that is dependent on multiple pattern recognition receptors. Finally, Mar1 is required for full virulence in two mouse models of systemic cryptococcosis.
Materials and methods
Strains, media, and growth conditions
Cryptococcus neoformans strains used in this study are listed in Table 1. Unless otherwise noted, all strains were generated in the C. neoformans var. grubii H99 background and maintained on YPD agar plates (2% yeast extract, 1% peptone, 2% dextrose). Strains created by crosses were co-incubated on MS mating media , followed by spore isolation by microdissection. Recombinant spores were identified by PCR and selectable marker resistance. To assess cell wall associated phenotypes, NaCl (1.5 M) and Congo red (0.5%) were added to YPD medium prior to autoclaving; caffeine (1 mg/ml) and calcofluor white (1 mg/ml) were filter sterilized and added to YPD after autoclaving. Alkaline pH plates were made by adding 150 mM HEPES buffer to YPD and adjusting the pH (pH 8.15) with NaOH prior to autoclaving. To induce and visualize capsule, strains were incubated in CO2-independent tissue culture medium (TC, Gibco) for 72 hours with shaking at 37°C, followed by staining with India Ink.
For cell wall staining, cell wall isolation, protein localization microscopy, FM4-64 staining, and in vitro co-culture experiments, overnight YPD cultures were diluted 1:10 in YPD liquid medium (at 30°C) or in TC medium (at 37°C) for 16–18 hours with shaking (150 rpm), unless otherwise noted. These methods were described previously .
All primers used in this study are listed in Table 2. All plasmids used in this study are listed in Table 3. The mar1ΔT-DNA strain was generated by Agrobacterium tumefaciens-mediated transformation (AMT) as described previously . C. neoformans targeted gene deletion strains were generated by homologous recombination to replace the entire open reading frame (ORF) with a dominant selectable marker. The deletion cassettes were created using overlap PCR as described previously [86,87] using the primers indicated in Table 2. Deletion cassettes were introduced into the indicated background strain by biolistic transformation . All deletion strains were confirmed by PCR and Southern blot; primers used to design probes for Southern blot can be found in Table 2.
The mar1Δ deletion strain (MAK1) was constructed by replacing the MAR1 ORF with the nourseothricin (NAT) cassette . The mar1Δ + MAR1 complemented strain (MAK11) was constructed by co-transformation of the WT MAR1 allele with the pJAF neomycin (NEO) resistance vector into the MAK1 background.
The GFP-Mar1 strain (SKE106) was constructed by transforming pSKE26 into the mar1Δ (MAK1) background. The pSKE26 plasmid contains an N-terminally GFP-tagged Mar1 protein under the control of the histone H3 promoter. A fragment consisting of the MAR1 open reading frame and ~ 500 bp of the 3’ UTR/terminator sequence was amplified from H99 genomic DNA using primer pair AA4894/AA4895. Using InFusion cloning (Clontech), this fragment was cloned in frame into the pCN50 backbone at a BamHI site at the end of GFP. Transformants were screened by wet colony morphology on pH 8 and GFP-Mar1 fusion was confirmed by PCR and western blot.
The mar1Δ + eGFP-RIM101 (MAK8) strain was constructed by crossing MAK1 with KS208. Recombinant spores were screened by epifluorescent microscopy and confirmed by PCR.
The MPK1-4FLAG-NEO strains (SKE94 and SKE96) were generated by transforming pSKE19 into the WT or mar1Δ (MAK1) background. The MPK1-4FLAG-NEO tagging construct was designed such that a C-terminal 4xFLAG epitope tag would homologously recombine into the 3’ end of the MPK1 locus. The pSKE19 plasmid was generated by In-Fusion cloning (Clontech) the following fragments into the pUC19 backbone: (1) ~500 bp of the 3’ end of MPK1 ORF, amplified from H99 genomic DNA using primer pair AA4829/AA4830; (2) 4xFLAG linked to the HOG1 terminator, amplified from pSG27  using primer pair AA4831/AA4832; (3) NEO resistance cassette amplified from pJAF using primer pair AA4264/AA4598; (4) ~ 1 kb 3’ MPK1 flank amplified from H99 genomic DNA using primer pair AA4599/AA4833. Transformants were screened by PCR and confirmed by Southern blot.
The mar1Δ mpk1Δ (SKE87) strain was constructed by replacing the MAR1 ORF with the NEO cassette in the mpk1Δ deletion background (KK3). Transformants were screened by dry colony morphology on pH 8 and PCR.
The FKS1-GFP-NEO strain (KMP13) was constructed by transforming pKP6 into the WT background. The FKS1-GFP-NEO tagging construct was designed to facilitate homologous recombination at the FKS1 locus. The pKP6 plasmid was created by In-Fusion cloning the following fragments into the pUC19 backbone: (1) ~ 1 kb of the 3’ end of the FKS1 ORF, amplified from H99 genomic DNA using primer pair AA4545/AA4362; (2) GFP amplified from pCN19 (Price 2008) using primer pair AA4364/AA4400; (3) FKS1 terminator (464 bp) amplified from H99 genomic DNA using primer pair AA4553/AA4426; (4) NEO resistance cassette amplified from pJAF using primer pair AA4264/AA1668; (5) 1 kb 3’ FKS1 flank amplified from H99 genomic DNA using primer pair AA4433/AA4546. Transformants were screened by PCR and epifluorescent microscopy and integration into the locus was confirmed by PCR. The mar1Δ FKS1-GFP-NEO strains (CLT1 and CLT2) were generated by replacing the MAR1 ORF with the NAT cassette in the KMP13 background. Transformants were screened by dry colony morphology on pH 8 and confirmed by PCR and Southern blot.
The mar1Δ cap59Δ (SKE60) strain was created by crossing MAK1 with CBN377. Recombinant spores were screened and confirmed by PCR.
The relative amount of capsule shedding in the cell supernatant was assayed as previously described [21,30]. Briefly, capsule induced cultures (incubated as described above) were incubated at 70°C for 15 minutes to denature enzymes, after which the cells were pelleted and the supernatant was sterile filtered. This conditioned medium was then run on a 0.6% agarose gel for 15 hours at 25 V, followed by transfer to a positively charged nylon membrane using Southern blotting methods. The membrane was air dried overnight, followed by blocking with 5% skim milk in Tris-Buffered Saline-Tween-20 (TBST). To detect capsule polysaccharide, blots were incubated with a mouse monoclonal anti-GXM antibody, MAb18B7 (1 μg/ml) [92,93] for 1 hour, washed 3x with TBST, and incubated with an anti-mouse horseradish peroxidase-conjugated secondary antibody (Jackson ImmunoResearch) for 1 hour. Blots were washed 3x with TBST and capsule polysaccharide was detected by enhanced chemiluminescence (ECL Prime Western blotting detection reagent; GE Healthcare).
Cell wall staining and flow cytometry
Prior to all cell wall staining, cells were pelleted and washed 1-2x with phosphate buffered saline (PBS). For quantification by microscopy, stained cells were imaged on a Zeiss Axio Imager.A1 fluorescence microscope equipped with an AxioCam MRm digital camera (60X objective). The same exposure time was used to image all strains with the same stains. The mean gray value (sum of gray values for all the pixels in a cell divided by the number of pixels that make up the cell) of at least 100 cells was calculated using ImageJ/Fiji [94,95]. Results are reported as mean fluorescence values +/- standard error of the means.
For flow cytometry, cells were fixed with 3.7% formaldehyde for 5 minutes at room temperature, followed by washing 2x with PBS. Eosin Y stained samples were fixed with 10 mM sodium azide for 10 minutes at room temperature, followed by washing 2x with PBS. A total of 107 cells/ml were stained and 106 cells/ml were submitted to the Duke Cancer Institute Flow Cytometry Shared Resource for analysis using a BD FACSCanto II flow cytometer. Data was analyzed using FlowJo v10.1 software (FlowJo, LLC). Relevant events were gated in the forward scatter/side scatter (FSC/SSC) plots and then represented as histograms with mean fluorescence intensity (MFI) on the x-axis and cell counts on the y-axis. Unstained cells and cells incubated with secondary antibodies alone were used as negative controls.
To visualize chitin, cells were stained with 100 μg/ml FITC-conjugated wheat germ agglutinin (WGA; Molecular Probes) for 35 minutes in the dark, followed by 25 μg/ml calcofluor white (CFW) for 10 minutes. Prior to analysis, cells were washed 2x and resuspended in PBS. For microscopy, WGA was imaged using a GFP filter and CFW was imaged using a DAPI filter. For flow cytometry, WGA cells were analyzed using a 488 nm laser and CFW cells were analyzed using a 405 nm laser.
To visualize chitosan, sodium azide fixed cells were washed 2x with McIlvaine’s buffer (0.2 M Na2HPO4, 0.1 M citric acid, pH 6.0), followed by staining with 300 μg/ml Eosin Y (EY) in McIlvaine’s buffer for 5 minutes at room temperature. Prior to analysis, cells were then washed 2x and resuspended in McIlvaine’s buffer. For microscopy cells were visualized using a GFP filter. For flow cytometry cells were analyzed using a 488 nm laser.
The MOPC-104E antibody (Sigma) was used to visualize α-glucan, as described previously [3,14]. Briefly cells were incubated with 400 ng/ml MOPC-104E primary antibody for 1 hour, washed 2x with PBS, and incubated with 4 μg/ml anti-mouse AlexaFluor 488 secondary antibody (Jackson ImmunoResearch) for 30 minutes in the dark. Cells were washed 2x and resuspended in PBS prior to analysis. For flow cytometry cells were analyzed using a 488 nm laser.
An Fc-Dectin-1 fusion protein was used to visualize β-glucan (a gift from Gordon Brown, University of Aberdeen) [96–99]. Cells were resuspended in FACS block (0.5% BSA, 5% HI-rabbit serum, 5 mM EDTA, 2 mM NaAzide in PBS) for 10 minutes, followed by incubation with 5 μg/ml Fc-Dectin-1 protein for 40 minutes on ice. Cells were washed 3x with PBS and resuspended in 3.75 μg/ml anti-human AlexaFluor 488 secondary antibody (Jackson ImmunoResearch) in FACS wash (0.5% BSA, 5 mM EDTA, 2 mM NaAzide in PBS) for 30 minutes on ice. Cells were washed 2x and resuspended in PBS prior to analysis. For flow cytometry cells were analyzed using a 488 nm laser.
Concanavalin A conjugated to AlexFluor 488 (ConA; Molecular Probes) was used to visualize mannoproteins. Cells were resuspended in 50 μg/ml ConA for 1 hour, then washed 2x and resuspended in PBS prior to analysis. For flow cytometry cells were analyzed using a 488 nm laser.
Differential interference microscopy (DIC) and fluorescent images were visualized with a Zeiss Axio Imager.A1 fluorescence microscope (60X or 100X objectives). Images were taken with an AxioCam MRm digital camera with ZEN Pro software (Zeiss). High-resolution fluorescent images were taken using a DeltaVision Elite deconvolution microscope equipped with a CoolSnap HQ2 high-resolution charge-coupled-device (CCD) camera. Images were processed using softWoRx software (GE). Images taken on both microscopes were additionally analyzed using ImageJ/Fiji software [94,95].
Cell wall isolation and analysis
Chitin and chitosan levels were quantified using a modified MBTH (3-methyl-benzothiazolinone hydrazine hydrochloride) method as previously described . Cell wall isolation and high performance anion-exchange chromatography with pulsed amperometric detection (HPAEC-PAD) were performed as previously described [16,33].
RNA extraction and real time PCR analysis
Cells from an overnight YPD culture were washed 1x with water, diluted to 107 cells/ml in YPD or TC medium in duplicate and incubated for 1.5 hours at 30°C (YPD) or 37°C (TC). Cultures were spun down and flash frozen on dry ice, followed by lyophilization. RNA was extracted using the RNeasy Plant Mini Kit (Qiagen) with optional on-column DNAse digestion. cDNA for real time-PCR (RT-PCR) was prepared using the AffinityScript cDNA synthesis kit (Agilent) with oligo(dT) primers. For RT-PCR, cDNA was diluted 1:3 in RNase-free water, added to IQ SYBR Green Supermix (Bio-Rad) per protocol instructions, and analyzed on an iCycler iQ Real-Time PCR Detection System (Bio-Rad). GPD1 was used as an internal control, and negative control samples without reverse transcriptase were included. All RT-PCR primers are listed in Table 2.
Protein extraction and western blot analysis
To assess Rim101 processing, overnight cultures were diluted to an optical density of 1 in 25 ml YPD pH 4 and pH 8. Cells were incubated for 1 hour, washed 1x with water, flash frozen on dry ice, and stored at– 80°C until cell harvesting.
For cell wall integrity pathway activation analysis, overnight YPD cultures were diluted to an optical density of 0.8 in 25 ml YPD in duplicate and incubated for 2.5 hours at 30°C. Cultures were then spun down and duplicates were resuspended in YPD or TC medium and incubated at 30°C (YPD) or 37°C (TC). After 3.5 hours, samples were taken, washed 1x with water, flash frozen on dry ice, and stored at -80°C until cell harvesting.
For protein extraction, cells were lysed by bead beating and the lysate was collected in 1.4 ml NP40 lysis buffer (6 mM Na2HPO4, 4 mM NaH2PO4, 1% Nonidet P-40, 150 mM NaCl, 2 mM EDTA, 1x protease inhibitors [Complete mini, EDTA-free; Roche], 1x phosphatase inhibitors [Phos-Stop; Roche], and 1 mM phenylmethylsulfonyl fluoride [PMSF]), as described previously . For Western blot analysis, samples were normalized by BCA assay (Thermo Scientific), diluted in 4x lithium dodecyl sulfate (LDS) loading sample buffer, and boiled for 5 minutes. Normalized protein was loaded on a NuPage 4–12% Bis-Tris gel (Invitrogen) and western blots were performed as previously described . To detect Gfp-Rim101, blots were incubated with an anti-GFP primary antibody (1/5,000 dilution; Roche) and an anti-mouse peroxidase-conjugated secondary antibody (1/25,000 dilution; Jackson Labs). To detect phosphorylated Mpk1, blots were incubated with a phospho-p44/42 MAPK (Thr202/Tyr204) rabbit polyclonal primary antibody (1/2,500 dilution; 4370 Cell Signaling Technology) and an anti-rabbit peroxidase-conjugated secondary antibody (1/50,000 dilution; Jackson Labs). Proteins were detected by enhanced chemiluminescence (ECL Prime Western blotting detection reagent; GE Healthcare).
FM4-64 staining and imaging
Cells from an overnight YPD (30°C) or TC (37°C) culture were normalized and stained with the lipophilic dye, FM4-64 (1/1000 dilution in the indicated medium, Molecular Probes) for 30 minutes shaking, after which the cells were pelleted and YPD or TC medium was refreshed for an additional 30 minutes at with shaking. For microscopy, cells were pelleted, washed 2x with PBS, and resuspended in PBS. FM4-64 was visualized using a Texas Red filter. The same exposure time was used for all images.
Acid and alkaline secretion analysis
Cells were incubated overnight in phosphate replete minimal medium (15 mM dextrose, 10 mM MgSO4, 13 mM glycine, 3 μM thymine, 0.4% KH2PO4). Cultures were then diluted to an OD of 0.9 in phosphate replete or phosphate deficient minimal medium (15 mM dextrose, 10 mM MgSO4, 13 mM glycine, 3 μM thymine, 0.4% KCl) and washed 1x in respective minimal media. 100 μl aliquots were plated in triplicate in a 96-well plate and incubated for 3 hours at 30°C with shaking at 150 rpm. Para-Nitrophenylphoshate (pNPP) solutions were prepared by dissolving a 5 mg pNPP substrate tablet (Thermo) in 50 mM sodium acetate (pH 5.2) for acid phosphatase testing or 1x diethanolamine substrate buffer (Thermo) for alkaline phosphatase testing. 100 μl of pNPP solution was added to each well and plates were incubated for 3 hours at 37°C with shaking at 150 rpm. Phosphatase activity was measured at an absorbance of 410 nm and adjusted for cell density, as determined by absorbance at 600 nm, over a time course of 3 hours.
Transmission electron microscopy
Overnight YPD and TC cultures were diluted to an OD of 0.5 and recovered in the same media for 4–6 hours. Cells were prepared as described previously [14,68,101]. Briefly, cells were spun down and washed 1x in pre-fixation mix (0.1 M sorbitol, 1 mM MgCl2, 1 mM CaCl2, 2% gluteraldehyde in 0.1 M PIPES, pH 6.8), followed by fixing in fresh pre-fixation mix overnight at 4°C. The next day, cells were washed 3x for 10-minute intervals in water. Next the cells were washed 3x in 2% KMnO4, and post-fixed for 45 minutes at room temperature in fresh 2% KMnO4. The cells were then washed repeatedly in water until no purple color was visible, and partially dehydrated for 10-minute intervals in increasing concentrations of ethanol (30%, 50%, 70%). Partially dehydrated samples were submitted to the Duke Shared Materials Instrumentation Facility (SMIF) for further processing, embedding, and sectioning as follows: Samples were rinsed thoroughly in PBS and post-fixed in 1% osmium tetroxide for 1 hour at room temperature. Samples were then stained with 0.5% uranyl acetate for 1 hour, further dehydrated in a series of graded ethanol (30%, 50%, 70%, 90%, 100%) and infiltrated overnight in resin. Samples were then embedded in resin and cured in a 55°C oven for 48 hours. The cured samples were thin sectioned with an ultramicrotome to approximately 60–90 nm. Thin-sections were mounted on copper grids and stained with uranyl acetate and lead citrate to enhance contrast. Grids were examined and digital images were taken on the FEI Tecnai G2 Twin transmission electron microscope with an Eagle digital camera.
Generation of bone marrow derived macrophages and dendritic cells
Bone marrow cells derived from female C57BL/6 mice purchased from Jackson Laboratories were used as WT controls for all experiments, unless otherwise noted. MyD88-/- and TLR2/4-/- bone marrow cells were a generous gift from Marcel Wüthrich at the University of Wisconsin-Madison. Bone marrow cells from Card9-/- mice were provided by Floyd Wormley. Dectin1-/- mice were a generous gift from Mari Shinohara at Duke University. C3H/HeOuJ TLR4 mutant mice (Stock no. 000659), C3H/HeJ control mice (Stock no. 000635), and TLR2-/- mice (Stock no. 004650, ) were purchased from Jackson Laboratories.
Murine bone marrow cells were isolated and prepared as previously described [16,103,104]. Briefly, femurs and tibias were isolated from mice and each bone was flushed with 5 to 10 ml cold PBS using a 27½ gauge needle. Red blood cells were lysed in 1x RBC lysis buffer (0.15 M NH4Cl, 1 mM NaHCO3, pH 7.4) and cells were resuspended in 1x Dulbecco’s modified Eagle’s medium (DMEM; + 4.5 g/L D-Glucose, + L-Glutamine, +110 mg/L sodium pyruvate) with 1 U/ml pencillin/streptomycin. Bone marrow cells were cryopreserved in 90% FBS/10% endotoxin-free DMSO at a concentration of 1 x 107 cells/ml and later thawed for use as previously described .
Fresh or frozen bone marrow cells were used to generate bone marrow derived macrophages (BMMs) or bone marrow derived dendritic cells (BMDCs). BMMs were differentiated in BMM medium (1x Dulbecco’s modified Eagle’s medium [DMEM; + 4.5 g/L D-Glucose, + L-Glutamine, +110 mg/L sodium pyruvate], 10% fetal bovine serum [FBS; non-heat inactivated], 1 U/ml penicillin/streptomycin) with 3 ng/ml recombinant mouse GM-CSF (rGM-CSF; R&D Systems or BioLegend) at a concentration of 2.5 x 105 cells/ml in 150 x 15 mm petri plates at 37°C with 5% CO2. The media was refreshed after 3–4 days and the cells were harvested on day 7 as previously described . BMMs were counted (by hemocytometer, with Trypan blue to discount dead cells), plated in BMM medium in 96-well plates at a concentration of 5 x 104 cells/well, and incubated at 37°C with 5% CO2 overnight prior to fungal co-culture experiments.
BMDCs were differentiated in BMDC medium (1x RPMI, 10% FBS [non-heat inactivated], 1 U/ml penicillin/streptomycin, 1x beta-mercaptoethanol) with 20 ng/ml rGM-CSF at a concentration of 5 x 106 cells/ml in 20 ml in 150 x 15 mm petri plates at 37°C with 5% CO2. After 3 days an additional 20 ml of BMDC medium with 20 ng/ml rGM-CSF was added to plates. After 6 days, 20 ml of culture supernatant was collected, centrifuged, resuspended in fresh BMDC medium with rGM-CSF and returned to the culture plate. BMDCs were harvested on day 10 as described previously  and BMDCs were counted, plated in BMDC medium in 96-well plates at a concentration of 5 x 104 cells/well, and incubated at 37°C with 5% CO2 overnight prior to fungal co-culture experiments.
In vitro fungal co-culture experiments
BMM and BMDC co-cultures with C. neoformans were performed as described previously . Briefly, C. neoformans cells were washed 2x with PBS, counted, and added to BMM or BMDC containing 96-well plates at a concentration of 5 x 105 fungal cells per well (10:1 C. neoformans cells:BMMs/BMDCs). Isolated cell wall material was added at a concentration of 10 mg/ml. Co-cultures were incubated for the indicated amount of time at 37°C with 5% CO2. Supernatants were collected and stored at -80°C until analysis. Secreted cytokines (TNF-α) were quantified in supernatants by enzyme-linked immunosorbent assay (ELISA; BioLegend). Data are represented as the average TNF-α values (pg/ml) for biological replicates; each fungal strain was tested a minimum of 3 times. BMM/BMDC only control wells, in which fresh media was added in lieu of fungi are included as negative controls. Ultrapure lipopolysaccharide (List Biolabs) and zymosan from S. cerevisiae (Sigma) were diluted to the indicated concentrations in BMM medium and used as positive controls.
As described previously , the cap59Δ mutation causes cell aggregation that makes quantification by hemocytometer inaccurate. As a result, these strains were normalized to 2 mg wet cell pellet/ml of medium, which was used previously for other mutants with similar mass/cell ratios and approximates the milligram-per-milliliter concentration used for standard co-culture experiments .
We used the murine inhalation model of Cryptococcosis to assess virulence . For each strain, 9–10 female C57BL/6 mice and 9–10 male and female BALB/c mice were used. Mice were anesthetized by isoflurane inhalation and intranasally inoculated with 1 x 105 fungal cells of the following strains: WT (H99), mar1Δ (MAK1), and mar1Δ + MAR1 (MAK11). Mice were monitored over the course of 40 days and sacrificed based on clinical endpoints that predict mortality. The statistical significance of difference between survival curves of mice infected with different strains was determined by log-rank test with Bonferroni correction (GraphPad Prism).
An additional 9–10 mice per strain were intranasally inoculated as described above for organ burden. Mice (5 per time point) were sacrificed on days 1 and 4 post inoculation. From each mouse 1 lung was harvested, weighed, and homogenized in cold PBS. Colony forming units (CFU) were calculated by quantitative culture and are represented as CFU/gram of tissue. For BALB/c post-mortem CFU analysis, lungs and brains were harvested from mice and homogenized in 1 ml PBS. Viable cells were calculated by quantitative culture and are represented as CFU/ml.
All animal experiments were performed in compliance with guidelines at Duke University, the University of Texas at San Antonio, and the American Veterinary Medical Association. All mice were anesthetized by isoflurane inhalation. Mice were sacrificed by CO2 with approved secondary methods of ensuring animal death. The Duke University Institutional Animal Care and Use Committee reviewed and approved the protocol (A138-17-06) used for animal experimentation in these studies. The specific projects were reviewed for congruence with this protocol, and approval was granted on 6/29/2015. Duke University maintains an animal program that is registered with the United States Department of Agriculture (Animal Welfare Act, Customer Number: 863), assured through the National Institutes of Health/Public Health Service (Assurance Number D16-00123 (A3195-01)), and accredited with AAALAC International (Accreditation Number: 363).
S1 Fig. Cell wall staining in additional conditions.
(A) WT cells incubated and stained as described in Fig 3. Bar, 10 μM. (B) Cell wall staining was assessed after incubation in YPD buffered to pH 8 or TC medium supplemented with 1x complete amino acids for 16–18 hours at 30°C with shaking. Cells were stained with FITC-conjugated WGA and imaged by fluorescent microscopy with the GFP filter.
S2 Fig. Mar1 is not regulated by or regulating the Rim pathway.
(A) MAR1 expression is induced in TC medium. WT cells were incubated for 1.5 hours in YPD (30°C) or TC (37°C), followed by RNA extraction and cDNA synthesis. Expression of MAR1 was determined by real-time PCR with fold change calculated relative to WT YPD levels and normalized to the expression of an internal control. Data represent means of results from 3 independent C. neoformans cultures and RNA extractions per condition. **, p = 0.0017 as determined by unpaired t-test. (B) Rim101 processing is intact in mar1Δ cells. WT and mar1Δ cells were incubated for 1 hour at the indicated pH, followed by western blotting using an α-GFP antibody.
S3 Fig. Mar1 is not a direct member of the cell wall integrity (CWI) pathway.
(A) CWI pathway signaling is intact in mar1Δ cells. WT and mar1Δ cells were incubated overnight in YPD and refreshed in YPD (30°C) or TC (37°C) for 3.5 hours, followed by western blotting using an α-phospho-Mpk1 antibody. Left panel is a representative blot image; Right panel is quantification of bands from 3 replicate experiments using ImageJ/Fiji software. (B) mar1Δ and mpk1Δ have combined effects on WGA staining. WT, mar1Δ, mpk1Δ, and mar1Δ mpk1Δ double mutant cells were incubated for 16–18 hours in YPD (30°C) or TC (30°C) followed by staining with WGA. Live cells were imaged by fluorescent microscopy and average fluorescence was quantified for at least 100 cells using ImageJ/Fiji software.
S4 Fig. Additional β-(1,3)-glucan synthase localization images.
Live cells were imaged using DeltaVision deconvolution fluorescent microscopy with the GFP filter. Images were deconvolved using softWoRx software. (A) Fks1-Gfp localization is similar in WT and mar1Δ mutant strains after incubation in YPD medium. Cells were incubated for 16–18 hours in YPD at 30°C prior to imaging. Bar, 10 μM. (B) Localization of Fks1-Gfp to the plasma membrane after incubation in TC media is decreased in an independent mar1Δ mutant. Cells were incubated for 16–18 in TC medium at 37°C prior to imaging. Bar, 10 μM.
S5 Fig. Macrophage activation by mar1Δ is independent of capsule.
Cultures of cap59Δ and mar1Δ cap59Δ were incubated for 16–18 hours in TC medium at 37°C. 2 mg/ml wet weight of each strain was co-cultured with BMMs for 6 hours, followed by quantification of TNF-α (pg/ml) in the supernatant by ELISA. Data represent 3 replicates from 3 independent experiments. **, p < 0.01 mar1Δ cap59Δ vs. cap59Δ as determined by one-way ANOVA with Tukey’s multiple comparisons test.
S6 Fig. C3H/HeJ and C3H/HeOuJ BMMs respond normally to control ligands.
BMMs were harvested from the indicated mouse strains and co-incubated with 10 ng/ml LPS or 10 μg/ml zymosan for 6 hours, followed by quantification of TNF-α (pg/ml) in the supernatant by ELISA. Data represent means of 3 replicates from 2 independent experiments (n = 6).
We acknowledge Jeanette Wagener and Louise Walker for performing the HPAEC-PAD analysis and Neil Gow for providing access to the Dionex HPAEC-PAD instrumentation. We thank Mike Cook and the Duke University Cancer Center Flow Cytometry Shared Resource for assistance with the flow cytometry. We also acknowledge Michelle Plue and the Duke University Shared Materials Institute Facility for performing the transmission electron microscopy. We thank Marcel Wüthrich for providing the MyD88-/-and TLR2/4-/- mice, and Mari Shinohara and Elizabeth Deerhake for providing the Dectin-1-/- mice.
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