Cellular morphology and morphogenesis are central features of fungal biology that contribute to the ability of fungi to cause disease in plants and mammals. In Candida albicans, one of the most common human fungal pathogens, the ability to transition between round, budding yeast and filamentous hyphae and pseudohyphae is positively correlated with its ability to cause both mucosal and invasive infections (Lopes and Lionakis, 2022; Sudbery, 2011). Conversely, establishment of the commensal state of C. albicans within the gastrointestinal tract is negatively correlated with filamentation (Witchley et al., 2019). Furthermore, the yeast form is thought to be required for dissemination within the blood stream while transition to hyphae contributes to tissue damage in both disseminated and mucosal disease (Saville et al., 2003; Meir et al., 2018). Consequently, C. albicans morphogenesis has been an intensively studied virulence trait (Villa et al., 2020).

To date, the extensive literature devoted to understanding the mechanisms of C. albicans morphogenesis has been almost entirely focused on the in vitro induction of the yeast-to-hyphae transition (Sudbery, 2011; Villa et al., 2020). Based on these studies, a wide range of genes and regulatory pathways have been found to modulate this transition in response to various inducing conditions in vitro. A significant knowledge gap is that it is not clear how many of these genes and pathways are also involved in C. albicans filamentation in vivo. Our group has developed an in vivo imaging assay that allows us to directly, and quantitatively, characterize C. albicans filamentation during mammalian infection (Wakade et al., 2021; Wakade et al., 2022b). Specifically, a 1:1 mixture of WT and mutant C. albicans strains labeled with different colored fluorescent proteins are inoculated into subcutaneous/epithelial stroma of a mouse ear. Using confocal microscopy, the relative yeast/filament ratio and the filament length of the two strains is readily determined for large numbers of cells, providing a statistically powerful assay of in vivo filamentation. Importantly, we can characterize both initiation of the morphological switch (ratio of yeast to filament) and maintenance of filamentous growth (filament length). Furthermore, the tissue that is infected in this approach is anatomically equivalent to the subepithelial stroma below mucosal surfaces where C. albicans colonizes humans. During initial disease, C. albicans undergoes yeast-to-hyphae transition within this compartment and thus our approach models filamentation in a tissue that is directly relevant to C. albicans pathogenesis.

Our initial investigations using this model have revealed significant differences between the in vitro and in vivo regulation of C. albicans filamentation (Wakade et al., 2021; Wakade et al., 2022a). For example, the cAMP-protein kinase A pathway is required for initiation of filamentation in almost all in vitro conditions; specifically, strains lacking the adenylyl cyclase gene (CYR1) or the PKA catalytic subunits (TPK1/2) remain in yeast phase under hyphae-inducing conditions (Cao et al., 2017). In contrast, cyr1∆∆ and tpk1∆∆ tpk2∆∆ mutants form hyphae in vivo to nearly the same extent as WT cells, indicating that this pathway plays a minor role in the regulating the switch between morphotypes in vivo (Wakade et al., 2022a). The length of the filaments formed by these mutants is shorter than WT (Wakade et al., 2022b). Consequently, the PKA pathway appears to switch from controlling initiation of hyphae formation in vitro to regulating maintenance of hyphal extension in vivo.

To further characterize the regulation of C. albicans filamentation in vivo, we screened a collection of 155 transcription factor (TF) deletion mutants (Homann et al., 2009) for those with either reduced hyphae/yeast ratios or filament length. We combined these studies with in vivo transcriptional profiling and genetic interaction analysis to provide the first model for the transcriptional network regulating C. albicans filamentation during infection of mammalian tissue. Although there are similarities between the TF networks operative in vitro and in vivo, we have identified TFs that affect filamentation in vivo but not in vitro. In addition, we provide evidence that the master regulator of C. albicans morphogenesis, Efg1, mainly functions to inhibit the function of Nrg1, a critical repressor of filamentation during filamentation within tissue.

C. albicans filamentation is one of the most extensively studied virulence traits of this important human fungal pathogen (Sudbery, 2011; Villa et al., 2020; Homann et al., 2009; Hirakawa et al., 2015). Prior to this work, systematic large-scale genetic analysis of filamentation was limited to in vitro induction of filamentation and very few in vivo studies had been reported (Homann et al., 2009; Uhl et al., 2003; Noble et al., 2010; O’Meara et al., 2015). The in vivo analysis of C. albicans filamentation using the direct inoculation of the subepithelial/subdermal tissue of mouse pinna followed by confocal microscopy reported herein has several features (Wakade et al., 2021; Wakade et al., 2022b; Wakade et al., 2022a). First, the method uses direct inoculation of the tissue and thus is not confounded by ability of mutants to colonize, disseminate, and/or invade target organs. Second, the subepithelial tissue is anatomically similar to the stroma below mucosal surfaces and is relevant to infection. Third, hypha-associated gene expression in infected ear tissue correlates well-infected kidney tissue (Wakade et al., 2022a), the main target of disseminated C. albicans infections in mice. Fourth, as we have demonstrated, the model allows the characterization of both filament initiation and elongation.

Efg1, Brg1, and Rob1 have the strongest phenotypes with respect to both filament initiation and hyphal elongation, although the efg1∆∆ mutant does not form filaments that can be assessed. These three TFs have similar phenotypes in vivo and in vitro (Figure 1); additionally, efg1∆∆ and brg1∆∆ mutants have reduced in vitro and in vivo filamentation across multiple clinical isolates (Wakade et al., 2021). The filamentation phenotypes of these three mutants have also been studied during GI colonization using a FISH-based assay (Witchley et al., 2019). During GI colonization, the efg1∆∆, brg1∆∆, and rob1∆∆ mutants were predominantly in yeast form suggesting they are important for filamentation in both niches. In contrast, the tec1∆∆ mutant formed near WT proportions of filaments during GI colonization (Witchley et al., 2019) while the proportion of filaments were reduced in the ear infection model, indicating that Tec1 may play a more important role in filamentation in the submucosae compared to colonization. Finally, Witchley et al. observed that the ume6∆∆ mutant formed filaments to an extent similar to WT in the gut (Witchley et al., 2019). We also observed no difference in the proportion of filaments formed by the ume6∆∆ mutant relative to WT but that the filaments were much shorter. Witchley et al. did not measure filament length in the GI model but inspection of their micrographs suggests that ume6∆∆ filaments were comparable in length to WT (Witchley et al., 2019). Thus, Efg1, Brg1, and Rob1 are important regulators of filamentation in these two niches whereas the roles of other TFs appear to vary from niche to niche.

In vitro analysis of filamentation/hyphae formation has been performed using a wide range of induction conditions and specific genes are required for filamentation in specific conditions. For example, Homann et al. screened the TF library that we used here using solid agar plates and 15 different media (Homann et al., 2009). Deletion mutants of BRG1, EFG1, ROB1 showed reduced filamentation on the majority of media/conditions while deletion of the repressors TUP1 and NRG1 showed increased filamentation under all condition. Our in vivo results for filament initiation correlate well with these trends indicating that TFs with important roles in vitro also have important roles in vivo. The mutants with less severe phenotypes in vivo such as tec1∆∆, ndt80∆∆, and cph2∆∆, for example, also had phenotypes in vitro but these were not as consistent across the different conditions. We suggest that these auxiliary regulators respond to a more limited set of stimuli or function together with one or more additional TFs. The synergistic interactions of Efg1 and Brg1 with Tec1 is an example where two core TFs function with an auxiliary TF to, in the case of Efg1-Tec1, interdependently regulate other auxiliary regulators (Figure 6D). Under some in vitro conditions, TFs such as Tec1 and Cph2 have very strong filamentation phenotypes (Homann et al., 2009; Lane et al., 2001) and we suggest that this may occur because those in vitro conditions generate a very specific filamentation signal that is highly dependent on a specific TF(s). In vivo, on the other hand, this signal is but one of many and therefore has a modest effect on filamentation process.

In the case of repressors of filamentation, a similar pattern arises where Tup1 and Nrg1 repress filamentation under nearly all in vitro conditions; in vivo, tup1∆∆ and nrg1∆∆ mutants were the only two with increased filamentation relative to WT. In vitro, 19 TF mutants have increased filamentation and, consequently, the largest discrepancy between in vitro and in vivo regulators is within the set of repressors (Homann et al., 2009). Of these, Nrg1, Tup1, and Ssn6 have been characterized as bona fide transcriptional repressors (Murad et al., 2001; Braun and Johnson, 1997; García-Sánchez et al., 2005) while the other TF mutants with increased filamentation are more likely to indirectly cause increased filamentation by altering the physiology of the cell such that its filamentation is a compensatory response. Although our assay may have reduced sensitivity for repressors, it is also possible that the in vivo environment is such a strong inducer of filamentation that mild to moderate repressor phenotypes are masked in this setting.

Nrg1 and Tup1 are widely described as functioning together to repress filamentation (García-Sánchez et al., 2005). Tup1 lacks a DNA-binding domain while Nrg1 does bind specific NRE-binding sites. However, while transcriptional profiling of tup1∆∆ and nrg1∆∆ mutants during filamentation showed significant overlap between the two regulons (García-Sánchez et al., 2005), distinctions in the differentially expressed genes were apparent as well. Furthermore, constitutive expression of NRG1 in the TET-off system blocks filamentation under in vitro inducing conditions (Saville et al., 2003), while expression of TUP1 from the same promoter does not (Ruben et al., 2020). These observations indicate that Tup1 and Nrg1 have genetically separable functions. Our data further support and extend this conclusion. The increased filamentation induced by deletion of TUP1 is dependent upon Efg1 (Figure 8A), under in vitro and in vivo conditions. In contrast, filamentation of the nrg1∆∆ mutant is not dependent on Efg1 (Figure 8D). The differential Efg1 dependence of Tup1 and Nrg1 is hard to reconcile with a model in which these two repressors function as a complex. Ssn6 and Tup1 are also a well-characterized co-repressor pair that is conserved in eukaryotic cells and have been proposed to function with Nrg1 as complex (García-Sánchez et al., 2005). However, we observed no phenotype for the ssn6∆∆ mutant in vivo; in vitro, the increased filamentation for ssn6∆∆ is not consistently observed. Taken together, our data and those in the literature suggest that Tup1 and Nrg1 play distinct roles in the repression of filamentation and that the mechanisms mediating relief of their repression are also distinct.

A model for the interactions of Tup1, Efg1, and Nrg1 that is consistent with our data as well with literature data is as follows. Tup1 lacks a DNA-binding domain and its deletion mutant has effects on gene expression that overlap with Nrg1 based on previously reported microarray studies (García-Sánchez et al., 2005). We, therefore, propose that Tup1 may bind to Efg1 rather than to Nrg1 and in doing so inhibit Efg1 function (Figure 10). This would explain the overlapping expression profiles of Tup1 and Nrg1 and the differential genetic interactions between Tup1-Efg1 and Nrg1-Tup1. To date, no CHiP analysis of either Tup1 or Nrg1 has been reported during in vitro filamentation. Therefore, it is not possible to definitively identify shared direct targets of Tup1 and Nrg1. The overlapping expression profiles are explained by our proposed model which is also consistent with genetic interaction data we report herein. Accordingly, future detailed biochemical and binding studies of the Tup1-Nrg1-Efg1 axis are likely to provide interesting new insights into the molecular mechanisms of filamentation regulation.

Figure 10

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The genetic interaction analysis between the core positive and negative regulators of filamentation uncovered the surprising result that Efg1 is dispensable for the expression of hyphae-associated genes following relief of Nrg1 repression. Each of the Efg1 bound genes that were restored to WT expression by deletion of NRG1 in the efg1∆∆ mutant also contained an NRE (to our knowledge, no comprehensive analysis of Nrg1 binding has been reported). These data suggest that Efg1 functions, at some level, to mediate the relief of Nrg1. Since Efg1 and Nrg1 bind some of the same targets, a potential mechanism would be for Efg1 to directly displace Nrg1 from the promoters of hypha-associated genes and, thereby, set the stage for TFs such as Rob1, Brg1, and likely others to bind and directly activate gene expression. Other mechanisms are possible and, regardless of that specific mechanism, our findings indicate that during hypha-initiation Efg1 indirectly regulates the expression of hyphae-specific genes. Overall, these genetic interaction studies indicate that much remains to be learned about the molecular mechanism by which hyphal formation is initiated in C. albicans.

Based on our genetic screens and genetic interaction studies, we have formulated a genetic circuit for the positive and negative regulation of filament initiation in vivo (Figure 10). This circuit incorporates previously established concepts and functions derived from in vitro studies as well as the data from the current study. As outlined above, the most distinct aspect of this model is that Efg1 plays an indirect role in regulating the canonical set of hyphae-specific genes by mediating the relief of Nrg1 suppression, leaving the direct activation of these genes to other TFs. Although we have largely limited our in vivo transcriptional studies to the core TFs involved in filament initiation, we suspect that at least some of the auxiliary regulators also contribute to the regulation of hyphae-specific genes as we propose for Tec1.

In addition to providing new insights into the initiation of filament formation, we have generated the first large-scale data set of TFs that are required for filament elongation (Lu et al., 2014). A number of TF mutants selectively affect the elongation phase of filamentation in vivo and at least five of these do not affect elongation in vitro. Hms1 and Lys14 are two TFs that regulate hyphal elongation in vivo but have few or no other in vitro phenotypes (Pérez et al., 2013; Shapiro et al., 2012). Perez et al. found that both Hms1 and Lys14 are required for systemic infection, suggesting that reduced elongation may contribute to virulence (Pérez et al., 2013). Our in vivo transcriptional profiling of the HMS1 and LYS14 mutants suggest that they regulate a distinct set of genes relative to the core TFs involved in filament initiation. Future studies will be required to understand the relationship between reduced filament elongation in vivo and virulence.

At least two distinct mechanisms could contribute to the reduced elongation phenotype for a given TF mutant. First, TFs could specifically, and directly, regulate the expression of genes that are uniquely required for hyphal elongation. A prime candidate for this class of TF is Ume6 since it is expressed only during hyphal formation and is required for this process in vitro and in vivo (Banerjee et al., 2008). Second, TFs that have general effects on growth under the conditions of hyphae formation would be expected to reduce the rate or limit elongation. We did not do time course analysis in this work and, therefore, it is possible that some of the mutants with reduced apparent filament length would eventually match WT lengths after additional time. From our set of mutants with reduced length, the sef1∆∆ mutant is potential candidate for this class of TF. Sef1 is required for C. albicans to respond to the low iron state established by the host during infection. Deletion of SEF1 leads to reduced growth under low iron conditions (Chen et al., 2011) and, therefore, the reduced cellular iron levels in the sef1∆∆ mutant may reduce filament length simply by limiting the growth of the hyphae. Other mechanisms such as septin regulation, Spitzenkorper assembly, activation, and maintenance of polarized growth factors are all undoubtedly active (Sudbery, 2011). Additional studies will be required to dissect the molecular mechanisms by which hyphae elongation-related TFs function during this process.

Ume6 plays an interesting role in that its phenotype is limited to filament elongation but it appears to share targets with hyphae-initiating TFs, particularly with respect to canonical hyphae-associated genes. As such, it seems to function like a transitional TF that bridges the initiation and elongation phases. It is likely that Rob1, Brg1, and other TFs with effects on both initiation and elongation may also bridge the two phases. Indeed, Nrg1 and Brg1 form a negative feedback loop during in vitro filamentation that is thought to maintain hyphal elongation (Lu et al., 2012; Cleary et al., 2012). Additional work will be required to develop a deeper understanding of the transcriptional regulation of the elongation phase and its key targets, particularly since mutants with selective defects in this process have decreased infectivity and virulence.

It is important to consider specific limitations of our approach. First, our transcriptional analyses are based on a focused set of genes known to be regulated by different environmental condition based on in vitro experiments. Our previous reported analysis of these gene has shown that the expression of these genes in kidney, ear, and in our in vitro conditions is reasonably well correlated. However, we may be missing key targets for in vivo filamentation since no genome-wide analysis of gene expression has been performed in the ear. This also means the proportion of genes regulated by a given TF is the proportion of our set of genes and we cannot make conclusions about the totality of genes regulated by a given TF.

Second, we selected a single in vitro filamentation condition to which we compare our in vivo data. There are a wide range of conditions that induce hyphae formation in C. albicans. Our selection was based on two primary considerations: (1) RPMI with 10% serum at 37°C is somewhat host-like and widely used in the field for this reason; (2) the extent of filament formation by our WT strain at 4 hr induction matches the extent of filamentation we observed after 24 hr in vivo; (3) as mentioned above the expression profiles in this condition were reasonably well correlated. However, it is almost certain that if other in vitro conditions were used as a comparator then a different set of results and conclusions would have resulted.

Third, some of our genetic interaction analysis was based on complex haploinsufficiency using double heterozygous mutants. We used this approach because the homozygous mutants of the core regulators have very low levels of filamentation; therefore, the dynamic range available to assay a double homozygous mutant would be unlikely to provide a statistically interpretable result. The limitation is that the double heterozygotes may not be sensitive enough to reveal subtle interactions; as such, this approach is quite specific but is likely to miss subtle genetic interactions. In the case of the combination of positive and negative regulators, these concerns were not present and, therefore, we used double homozygous mutants.

Finally, it is important to consider the generalizability of these data and conclusions to other filament-inducing conditions and, most importantly, other host niches. As discussed above, it is becoming emphatically clear that the regulatory networks governing C. albicans filamentation vary from condition to condition and host niche to host niche. For example, the most widely recognized master transcriptional regulator of filamentation, Efg1, is dispensable for this process when the cells are embedded in agar and incubated at 25°C or under hypoxic conditions. From our own work, we have reported that the brg1∆∆ deletion mutant forms robust filaments in oral tissue but has greatly reduced filamentation in the ear model. Our focus on filamentation within the sub-stromal tissue of the ear epithelium is likely to yield results that have both similarities and differences to filamentation at other sites and under other conditions; however, these limitations are true for any study of C. albicans filamentation. We assert that the complexity of this key C. albicans virulence trait is worthy of additional study and, therein, likely to yield additional insights.

In summary, our systematic genetic analysis of the transcriptional regulation of C. albicans filamentation has both confirmed the importance of key regulators identified in vitro and expanded our understanding of the process to include details and concepts that could not have been inferred from in vitro studies alone. The latter emphasizes the utility of studying virulence-associated traits during infection as well as in vitro.

All data generated or analyzed during this study are included in the manuscript, supporting files, and source data files for Fig. 1, 2, and 8 are provided. Both the raw and processed gene expression data generated by Nanostring are provided in supplementary files 3,4, 5 and 6. No sequencing data was generated in this study.

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Author details

1. Rohan S Wakade

   Department of Pediatrics, Carver College of Medicine, University of Iowa, Iowa City, United States

   Contribution

   Data curation, Formal analysis, Investigation, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

2. Laura C Ristow

   Department of Pediatrics, Carver College of Medicine, University of Iowa, Iowa City, United States

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

3. Melanie Wellington

   Department of Pediatrics, Carver College of Medicine, University of Iowa, Iowa City, United States

   Contribution

   Conceptualization, Formal analysis, Supervision, Methodology, Writing – review and editing

   Competing interests

   No competing interests declared

4. Damian J Krysan

   1. Department of Pediatrics, Carver College of Medicine, University of Iowa, Iowa City, United States
   2. Departments of Microbiology and Immunology, Carver College of Medicine, University of Iowa, Iowa City, United States
   3. Molecular Physiology and Biophysics, Carver College of Medicine, University of Iowa, Iowa City, United States

   Contribution

   Conceptualization, Formal analysis, Supervision, Funding acquisition, Writing – original draft, Writing – review and editing

   For correspondence

   damian-krysan@uiowa.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-6330-3365

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