The transition to host CO₂ concentrations affects a wide range of cellular processes in C. neoformans

To identify genes with altered fitness at host concentrations of CO₂ (5%), a moderate throughput competitive fitness assay that we previously reported [12] was used to screen 4,692 strains from the Madhani C. neoformans deletion library; all deletion mutants were in the CO₂-tolerant KN99α strain background (S1 Table). Each mutant was inoculated into the well of a 96-well plate with an approximately equal cell concentration of mNeonGreen-labeled reference strain H99. The strains were incubated in RPMI buffered with MOPS in ambient air or 5% CO₂ at 37 °C for 24 h. The ratios of the two strains were quantified using flow cytometry, and the competitive fitness calculated as described in Materials and methods. Mutants with statistically significant (three replicates, P < 0.05 Student t test) competitive fitness changes of ±0.2 were considered to have altered fitness.

A total of 301 genes met these criteria with 245 mutants showing reduced fitness and 56 mutants displaying increased fitness (Fig 1A and S1 Table). We previously identified multiple kinase mutants in the Bahn Library that have altered fitness in 5% CO₂; importantly, a subset of these were also in the Madhani Library and showed the same fitness phenotypes [12], including mutants of MPK1, BCK1, MKK2, PKA1, ARK1, PKA1, CBK1, FBP26, YPK1, and CEX1. In addition, recently, it has been reported that plating errors in the distribution of the Madhani deletion set have occurred, leading to obvious problems with reproducibility. Our group has previously reported screening a subset of the Madhani library (obtained independently) under different conditions (yeast peptone 2% dextrose (YPD), 10% CO₂ on solid agar plates, [13]); a total of 25 mutants identified by Chadwick and colleagues were also identified in this work [13]. Finally, we have also independently constructed mutants for further study and confirmed those phenotypes. A summary of these corroboration results is provided in S2 Table. While large-scale screening experiments have inherent false positive and negative rates, our follow-up data indicate that the screening results are reasonably robust within limitations expected for a genome-wide mutant screen.

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Fig 1. Genome-wide deletion collection screening revealed a wide range of cellular processes involved in response to CO₂.

A. Volcano plot of the average competitive fitness score and significance of three biological replicates for each mutant in 5% CO₂ conditions relative to ambient air in the Madhani deletion collection. Competitive fitness was determined after combining a 1:1 ratio of mNeonGreen-labeled H99 and unlabeled mutant cells in RPMI 1,640 medium with 165 mM MOPS, pH 7, and incubating at 37 °C in ambient air or 5% CO₂ for 24 h. Cell populations were characterized by flow cytometry, and the percentage of mNeonGreen-negative cells in 5% CO₂ was divided by those in ambient conditions to calculate competitive fitness. Colored data points indicate values that were below 0.8 (red) or above 1.2 (blue) and statistically significant by Student t test (P < 0.05). Biological-process GO term analysis for the genes identified in (A) at 24 h are depicted in B (significantly reduce CO₂ fitness) and D (significantly increase CO₂ fitness). C. Cell lysates from H99 cells exposed to ambient air or 5% CO₂ for 2 h were analyzed by western blot for Hog1 expression and phosphorylation status. Results are representative of three biological replicates. E. Venn diagram comparing mutants with reduced CO₂ fitness (identified in A) to mutants with reduced fitness in a mouse model of cryptococcosis reported in [14]. E. Venn diagram comparing mutants with reduced fitness at 37 °C to mutants with reduced fitness in a mouse model of cryptococcosis reported in [14]. The data underlying this Figure can be found in S1 Table and S1 Data.

https://doi.org/10.1371/journal.pbio.3003561.g001

Biological process GO term analysis of the set with reduced fitness provided some insight into the overall functions involved in CO₂ tolerance (Fig 1B). Response to the detergent sodium dodecyl sulfate (SDS) and salt were the two terms with the lowest adjusted p values (Benjamini-Hochberg). SDS interferes with membrane function and is consistent with our previous findings that CO₂ leads to remodeling of the plasma membrane and that drugs targeting sterols (fluconazole) and sphingolipid biosynthesis (myriocin) are more active against C. neoformans at host CO₂ concentration than in ambient air [8].

The response to salt appears to be driven by a large set of protein kinases with pleomorphic phenotypes [14] that include sensitivity to salt (CCK1, IPK1, CBK1, YPK1, SNF1, TPS2). If the host CO₂ concentrations directly triggered a stress response that overlaps with salt stress, then it may increase activity of the Hyperosmolar Glycerol Pathway (HOG) pathway, an important part of the cellular response to elevated salt [15]. Our previous screen of a protein kinase library [12], however, did not find that mutants of the HOG1 pathway were less fit in CO₂ (in the current screen the hog1∆ mutant showed modestly reduced fitness 0.78, S1 Table). Furthermore, transition to high CO₂ did not change the phosphorylation status of Hog1 (Fig 1C). Therefore, it seems sensitivity to salt is an indirect correlation with CO₂ sensitivity. Finally, genes involved in response to oxidative stress were also enriched in those required for CO₂ fitness.

The set of gene mutations that increased CO₂ fitness is also enriched for response to salt along with response to pH and multivesicular body (MVB) formation (Fig 1D). The presence of the latter two groups of genes is likely related to the fact that the Rim101 pathway is maladaptive during CO₂ stress [12] and that some genes that contribute to the MVB are involved in Rim101 processing [16]. Interestingly, the set of genes required for full CO₂ fitness was enriched for those that are also annotated to affect the “big three” virulence traits [5,6] of capsule formation, melanin metabolism, and response to heat (Fig 1B). However, the set of genes whose loss of function increases fitness in 5% CO₂ was also enriched for those with effects on capsule, melaninization, and salt stress (Fig 1D). The connection between genes that affect CO₂ fitness and the “big three” virulence traits is likely due to the fact that all of these phenotypes are indirectly affected by many physiologic processes. Of these, capsule formation is the trait most likely to share physiologically direct connections to CO₂ fitness because it is a well-characterized inducer of capsule formation in vivo [17].

Finally, we compared the set of mutants with reduced CO₂ fitness to the set of mutants with reduced fitness in a mouse model of cryptococcosis as recently reported by Boucher and colleagues [14]. The Venn diagram in Fig 1E shows that 138/245 (56%) mutants with reduced CO₂ fitness in vitro also have reduced fitness during infection. We do not know what level of CO₂ fitness under these in vitro conditions is required to maintain in vivo fitness. However, we took advantage of the large-scale phenotyping data from Boucher and colleagues [14] to compare the CO₂/in vivo fitness correlation with that between fitness at body temperature (37 °C) and in vivo fitness. The ability to grow at host body temperature is one of the three well-established C. neoformans virulence traits. Interestingly, the overlap of mutants with reduced fitness at 37 °C with in vivo fitness was remarkably similar (152/272, 56%) to that for CO₂ fitness (Fig 1F). These comparisons suggest that both CO₂ and host body temperature tolerance are virulence traits that display variable penetrance rather than virulence factors that show a binary, present/absent, distribution. Further, these traits are likely to interact with each other as well as with additional virulence traits to determine the overall in vivo fitness of a given mutant or strain.

The RIM101 and PKA pathways negatively regulate CO₂ tolerance in an independent manner

The set of mutants associated with increased CO₂ fitness was enriched for those involved in response to alkaline pH (Fig 1D). In addition to re-identifying the rim101∆ mutant, deletion mutants of components of the pathway required for proteolytic processing (Rim13 and Rim20), and hence activation, of Rim101 were also found to have increased fitness in 5% CO₂. Activation of the Rim101 pathway has been linked to Endosomal Sorting Complex Required for Transport (ESCRT) complex function in model yeasts [18]. Furthermore, the Kronstad [19] and Alspaugh [16] labs have shown that the ESCRT complex is involved in Rim101 activation in C. neoformans as well. In keeping with these observations, deletion mutants of five genes with homology to ESCRT complex components (VPS22/23/25/28&36) showed increased fitness in CO₂.

In principle, the increased fitness of mutants related to tolerance of elevated pH could be due to pH-related functions or due to other functions with pH response as a common but distinct function. Therefore, we compared the CO₂ phenotypes of the ESCRT complex mutants to non-ESCRT complex VPS genes (Fig 2A) and re-tested them on YPD plates (S1 Fig); the CO₂ phenotypes were consistent between the two conditions for all strains. All five ESCRT complex mutants were more fit in CO₂ than H99 (VPS22/23/25/28&36) while non-ESCRT VPS mutants either had reduced CO₂ fitness (VPS1 and VPS31) or no phenotype (VPS29/35/&41); the increased fitness of the vps29∆ mutant is statistically significant but not likely to be large enough to be biologically relevant. Regardless of their CO₂ tolerance phenotype, all VPS mutants except for the vps29∆ mutant showed reduced growth on YPD adjusted to pH 8 (Fig 2B). Since both CO₂-tolerant and -intolerant VPS mutants are generally sensitive to pH 8, it appears that the CO₂ fitness phenotypes are not directly related to an impaired response to elevated pH but rather due to other functions of the ESCRT/Rim101 pathway or VPS genes.

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Fig 2. The RIM101 and PKA pathways negatively regulate CO₂ tolerance in an independent manner.

A. Competitive fitness was determined for the indicated vacuolar protein sorting (VPS) mutants after combining a 1:1 ratio of mNeonGreen-labeled H99 and unlabeled mutant cells in RPMI 1,640 medium with 165 mM MOPS at pH 7 and incubating at 37 °C in ambient air or 5% CO₂ for 24 h. Cell populations were characterized by flow cytometry, and the percentage of mNeonGreen-negative cells in 5% CO₂ was divided by those in ambient conditions to calculate competitive fitness. Bars represent the average of three biological replicates with error bars indicating SD. Asterisks indicate a significant difference relative to H99 as determined by Student t test (P < 0.05). B. Overnight cultures of rim101∆ and vps mutants were washed, and concentration determined by OD₆₀₀ reading. Strains were diluted to a starting OD₆₀₀:1, and serial 10-fold dilutions were performed. Three µL per replicate were spotted onto standard YPD agar plates, or YPD buffered with 150 mM HEPES and adjusted to pH 8. Plates were incubated at 37 °C in ambient air for 48 h before images were acquired. Images are representative of three biological replicates. Competitive fitness of pkr1∆ and pka1∆ mutants (C) and rim101 mutants (D) in 5% CO₂ as described in (A). The data underlying this Figure can be found in S1 Table and S1 Data.

https://doi.org/10.1371/journal.pbio.3003561.g002

O’Meara and colleagues have demonstrated that some functions of the Rim101 pathway are regulated by the Protein Kinase A (PKA) pathway [20]. Previously, we showed that, like loss of function mutants in the RIM101 pathway, deletion of PKA1 increases CO₂ fitness [12]; an observation we re-confirmed in this screen (Fig 2C). The PKA pathway is negatively regulated by Pkr1 and, consistent with that relationship, the pkr1∆ mutant is less fit in CO₂ (Fig 2C). Therefore, we tested whether the PKA pathway directly regulates Rim101 during CO₂ stress. To do so, we took advantage of a strain constructed by O’Meara and colleagues in which the only source of RIM101 is an allele in which the PKA phosphosite (Ser773) was mutated to alanine (rim101^Ser773Ala, [20]). The rim101^Ser773Ala mutant did not show a significant difference from WT in fitness under 5% CO₂ (Fig 2D). Therefore, both the PKA and RIM101 pathways negatively affect CO₂ tolerance, but they do so in a manner independent of each other.

Regulation of membrane homeostasis through both the TOR and UPR pathways is required for CO₂ tolerance

The TOR pathway is also a key positive regulator of CO₂ tolerance, and prior work has shown that it regulates remodeling of the plasma membrane asymmetry in response to elevated CO₂ concentrations [12]. Further validating the importance of the TOR pathway to CO₂ tolerance, two putative components of the TOR complex, SIN1 and a protein homologous to Avo2 (CNAG_02727), as well as SIT4, a phosphatase that appears to function downstream of TOR in S. cerevisiae [21], were among the mutants with reduced fitness (S1 Table). A key function of the Tor-Ypk1 axis in model yeast is the regulation of sphingolipid biosynthesis [22]. In S. cerevisiae, Ypk1 phosphorylates and activates the serine-palmityl transferase (SPT, [23]) which regulates the first committed step of sphingolipid biosynthesis (Fig 3A). As previously reported, the potency of the SPT inhibitor, myriocin, against C. neoformans is increased in 5% CO₂, indicating sphingolipid synthesis is required for CO₂ tolerance [8,12]. In addition, Lee and colleagues have previously demonstrated that Ypk1 is required for sphingolipid biosynthesis in C. neoformans [23].

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Fig 3. Regulation of membrane homeostasis through both the TOR and UPR pathways is required for CO₂ tolerance.

A. Schematic representation of the early steps in C. neoformans sphingolipid biosynthesis pathway. Genes required for CO₂ fitness are indicated in red. Created in BioRender. Ristow, L. (2026) https://BioRender.com/jgg870j. B. Competitive fitness was determined for glucosylceramide synthase mutant gcs1∆ after combining a 1:1 ratio of mNeonGreen-labeled H99 and unlabeled mutant cells in RPMI 1,640 medium with 165 mM MOPS at pH 7 and incubating at 37 °C in ambient air or 5% CO₂ for 24 h. Cell populations were characterized by flow cytometry, and the percentage of mNeonGreen-negative cells in 5% CO₂ was divided by those in ambient conditions to calculate competitive fitness. Bars represent the average of three biological replicates with error bars indicating SD. Asterisks indicate a significant difference relative to H99 as determined by Student t test (P < 0.05). C. Cells from overnight cultures of H99 were spread on solid RPMI agar plates buffered with 165 mM MOPS to pH 7. Sterile disks were placed on plates, and 100 µg of aureobasidin A was added to each disk. Plates were incubated at 37 °C in ambient air or 5% CO₂ for 48 h before images were acquired. Images are representative of three biological replicates. D. Competitive fitness of the unfolded protein response (UPR) mutants ire1∆ and hxl1∆ in 5% CO₂ as described in (B). The data underlying this Figure can be found in S1 Table.

https://doi.org/10.1371/journal.pbio.3003561.g003

Downstream of SPT (Fig 3A), the sphingolipid biosynthesis pathway has two branches: glucosylceramide (GC) and inositol-phosphoryl-ceramide (IPC). We identified the gcs1∆ mutant as having reduced CO₂ fitness, demonstrating the importance of the GC branch in CO₂ tolerance (Fig 3B). IPC synthase (Ipc1) is an essential gene [24] and, therefore, to determine if the IPC branch is also required for CO₂ tolerance, we compared the potency of the specific IPC inhibitor, aureobasidin A in ambient and 5% CO₂. Indeed, the zone of clearance induced by aureobasidin A is increased at 5% CO₂ relative to ambient air (Fig 3C). These data firmly establish that an essential function of the Tor-Ypk1 axis during CO₂ stress is to maintain sphingolipid homeostasis and that both major sphingolipid classes contribute to CO₂ tolerance.

Membrane homeostasis is also critically dependent on the function of the endoplasmic reticulum (ER) for the synthesis of lipids such as sterols, for the delivery to membrane proteins, and for the generation of the cell wall which buffers the cells against the effects of membrane-directed stressors. The unfolded-protein response (UPR) pathway functions to maintain ER homeostasis in the presence of a variety of stressors that affect its function [26]. The two key proteins in this pathway are the kinase IRE1 and the downstream transcription factor HXL1 (aka BZP1). Mutants lacking either component of the UPR pathway have reduced fitness in 5% CO₂ (Fig 3D). These data contribute to the growing body of evidence that host CO₂ exerts a significant stress on the membranes of C. neoformans and, consequently, key mediators of membrane biosynthesis and membrane stress response pathways are critical for the ability of C. neoformans to replicate under those conditions.

Host CO₂ concentrations remodel C. neoformans carbon metabolism

Through its conversion to bicarbonate, CO₂ is an essential substrate for many key reactions in central carbon metabolism [27]. At the same time, it is generated as a byproduct in other reactions of central carbon metabolism such as those in the tricarboxylic acid cycle in the mitochondria. Twenty-five mutants involving central carbon metabolic processes were identified as being required for CO₂ fitness (Table 1). Sixteen of these genes are associated with mitochondrial functions. In addition, the deletion mutants of two non-essential genes involved in glycolysis (FBP26 and PFK1) have reduced fitness as do five genes associated with the synthesis of acetyl-CoA or CoASH (YHM2, CAB3, LSC1, POT1, and ALD4). Lastly, multiple genes in the kynurenine pathway, which generates nicotinamide, were also less fit in 5% CO₂.

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Table 1. Carbon metabolism genes required for fitness in host concentrations of CO₂.

https://doi.org/10.1371/journal.pbio.3003561.t001

To further explore the effect of host CO₂ concentrations on the metabolic state of C. neoformans, we compared the non-lipid metabolomic profile of C. neoformans incubated in buffered RPMI at ambient CO₂ to that from cells incubated in 5% CO₂ after 24 h at 37 °C. The abundance of 90 molecules differed between 5% CO₂ and ambient air (± 1 log₂, adjust P < 0.05, S3 Table). Strikingly, glucose; two glycolysis intermediates, phosphoenopyruvate (PEP) and dihydroxyacetone-3-phosphate (DHAP); and fructose (Fig 4A) are reduced, suggesting that exposure to 5% CO₂ depletes glucose and glycolytic intermediates.

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Fig 4. Host CO₂ concentrations remodel C. neoformans carbon metabolism.

The normalized area of each metabolite was determined by tandem mass spectrometry as described in materials and methods for H99 incubated in buffered RPMI in ambient air or 5% CO₂. Asterisks indicate statistically significant differences in abundance at 5% CO₂ relative to ambient air (Welch corrected, two-tailed t test, P < 0.05). A. Relative amounts of glycolytic intermediates. B. Relative amounts of UDP-Glucuronic Acid (UDP-GluA). C. Competitive fitness of the uxs1∆ and uge1∆ mutants relative to H99 was determined for indicated strains after combining a 1:1 ratio of mNeonGreen-labeled H99 and unlabeled mutant cells in RPMI 1,640 medium with 165 mM MOPS at pH 7 and incubating at 37 °C in ambient air or 5% CO₂ for 24 h. Cell populations were characterized by flow cytometry and the percentage of mNeonGreen-negative cells in 5% CO₂ was divided by those in ambient conditions to calculate competitive fitness. Bars represent the average of three biological replicates with error bars indicating SD. Asterisks indicate a significant difference relative to H99 as determined by Students t test (P < 0.05). D. H99 was incubated in buffered RPMI medium with standard glucose (0.2%) or the indicated amount of glucose supplementation at 37 °C at ambient or 5% CO₂. The optical density of the cultures was measured at 48 h. Growth was normalized to that of standard RPMI for each CO₂ concentration. The bars indicate mean of two independent replicates, and error bars indicate standard deviation. The differences between conditions were analyzed by One-way ANOVA with post hoc correction for multiple comparisons. The data underlying this Figure can be found in S1 and S3 Tables and S1 Data.

https://doi.org/10.1371/journal.pbio.3003561.g004

Host CO₂ is a strong inducer of the polysaccharide capsule in C. neoformans [17], and we have previously shown that genes involved in capsule biosynthesis are enriched in the set of genes upregulated by exposure of H99 to host levels of CO₂ [12]. The carbohydrates that make up the capsule are derived from glucose, suggesting that host CO₂ may divert glucose from glycolysis to capsule biosynthesis [28]. Indeed, the levels of a glucose-derived capsule biosynthesis intermediate, UDP-glucuronate (UDP-GlnA), are reduced 4-fold in 5% CO₂ relative to ambient CO₂ levels (Fig 4B), supporting the idea that flux through this pathway is increased. In the next step of capsule biosynthesis, UDP-GlnA is converted to UDP-xylose by UDP-xylose synthase (UXS1, [29]). Consistent with this function, the uxs1∆ mutant has reduced capsule thickness [30]. Deletion of UXS1 increases CO₂ fitness (Fig 4C), suggesting that interrupting capsule biosynthesis early in the pathway provides a growth advantage. In contrast, deletion of UGE1, the enzyme that epimerizes UPD-glucose to UDP-galactose, increases capsule thickness [31] and reduces CO₂ fitness (Fig 4C).

If this model is operative, we hypothesized that increasing glucose concentrations would improve growth of H99 in 5%CO₂. RPMI medium contains 0.2% glucose, and we previously reported that increasing glucose concentrations to 2% did not affect the growth of H99 in 5% CO₂ [9]; therefore, we supplemented the medium to 2%, 5%, and 10% and incubated H99 at 37 °C in either ambient or 5% CO₂ for 48 h. Cultures of H99 containing increased glucose that were incubated at ambient CO₂ reached a higher final cell density relative to standard RPMI; however, there was no apparent dose effect between 2%–10%, indicating that at glucose concentrations above 2% other nutrients were growth limiting (Fig 4D). In 5% CO₂, the effect of increasing glucose concentrations on the final cell density is blunted relative to ambient CO₂ (Fig 4D). A statistically significant effect on final cell density is not observed until the concentration of glucose is 10%.

These data support the model that at host CO₂ concentrations, glucose is being diverted to processes other than replication and that high glucose concentrations can saturate the non-replicative pathway which, in turn, leads to increased final culture density. In contrast, at ambient CO₂, 2% glucose is sufficient to render glucose a non-limiting nutrient. Taken together with our genetic and metabolic data, we propose that CO₂-induced capsule formation diverts glucose from glycolysis and other energy-generating processes, leading to the use of alternative carbon sources at host CO₂ concentration.

Protein catabolism and urea cycle activity is increased in response to host CO₂ concentrations

Metabolomic studies in other yeast have shown that depletion of glucose increases protein catabolism in order to mobilize amino acids as carbon sources [32]. As shown in Fig 5A, nine di- and tripeptides are increased at 5% CO₂, a strong indication that C. neoformans protein catabolism is operative under these conditions. In addition, the amounts of six amino acids (Leu, Phe, Tyr, Trp, His, and Met) are increased by ≥ 2-fold in cells grown in 5% CO₂ (S3 Table). Of the nine di- and tri-peptides identified, all contain ketogenic amino acids, and 7/9 peptides contain Leu and Lys which are exclusively ketogenic amino acids. Ketogenic amino acids can be further catabolized in the mitochondria to acetyl-CoA which then enters the TCA cycle as an alternative to acetyl-CoA derived from glycolysis-derived pyruvate [33]. Thus, it appears that C. neoformans mobilizes alternative carbon nutrients by protein catabolism to compensate for reduced amounts of glucose after shift to host levels of CO₂.

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Fig 5. Host CO₂ concentrations induce increased protein catabolism and urea cycle activity.

The normalized area of each metabolite was determined by tandem mass spectrometry as described in materials and methods for H99 incubated in buffered RPMI in ambient air or 5% CO₂. Asterisks indicate statistically significant differences in abundance at 5% CO₂ relative to ambient air (Welch corrected, two-tailed t test, P < 0.05). A. Relative abundance of the indicated di- and tripeptides in ambient and 5% CO₂. B. Competitive fitness of mutants involved in vacuolar ATPase function. Competitive fitness of the indicated VMA mutants relative to H99 was determined for indicated strains after combining a 1:1 ratio of mNeonGreen-labeled H99 and unlabeled mutant cells in RPMI 1,640 medium with 165 mM MOPS at pH 7 and incubating at 37 °C in ambient air or 5% CO₂ for 24 h. Cell populations were characterized by flow cytometry, and the percentage of mNeonGreen-negative cells in 5% CO₂ was divided by those in ambient conditions to calculate competitive fitness. Bars represent the average of three biological replicates with error bars indicating SD. Asterisks indicate a significant difference relative to H99 as determined by Students t test (P < 0.05). C. Schematic of the initial steps of the urea cycle in C. neoformans. Created in BioRender.com. Ristow, L. (2026) https://BioRender.com/3jm5mu3. Relative abundance of the urea cycle intermediates citrulline (D) and argininosuccinate (E) in ambient and 5% CO₂. The data underlying this Figure can be found in S1 and S3 Tables.

https://doi.org/10.1371/journal.pbio.3003561.g005

Protein catabolism occurs in the vacuole and is dependent upon vacuolar ATPase (V-ATPase) to generate the acidic environment necessary for the activation of vacuolar peptidases [34]. Multiple strains lacking genes coding for V-ATPase components or proteins involved in the assembly of the V-ATPase complex show reduced CO₂ fitness (Fig 5B), indicating that maintenance of vacuolar pH is critical for tolerance of high CO₂. Although V-ATPase mutants have pleomorphic effects on cells, the metabolomic data clearly indicate protein catabolism occurs during the initial phases of CO₂ stress. Accordingly, loss of V-ATPase function would likely interfere with efficient generation of ketogenic amino acids necessary to maintain mitochondrial respiration.

CO₂/HCO₃^- is essential for the cell, but high levels of CO₂ are stressful and can ultimately be toxic. As such, CO₂ is both generated and consumed by metabolic reactions in the cell. The urea cycle is one of the CO₂-consuming processes utilized by eukaryotic cells (Fig 5C). Ura2 is a carbamoyl phosphate synthase that catalyzes the rate-limiting condensation of CO₂ with ammonia which feeds into the urea cycle. Previous work by our group found that deletion of URA2 reduces CO₂ fitness [11] and our screen confirmed this result (Table 1). Carbamoyl-phosphate enters the urea cycle by reacting with ornithine to generate citrulline which, in turn, is converted to argininosuccinic acid (Fig 5C). Both the citrulline and argininosuccinic acid levels are elevated at 5% CO₂ relative to ambient levels (Fig 5D and 5E), suggesting increased conversion of CO₂ to urea cycle intermediates relative to ambient CO₂. Supporting this conclusion, in S. cerevisiae, urea cycle activity is enhanced under increased CO₂ concentrations to deplete excess CO₂/HCO₃ [35]. Ura2 is also required for the synthesis of uridine/uracil and deletion mutants are auxotrophic for those nutrients. However, Chadwick and colleagues found that the reduced fitness of the ura2∆ mutant in 5% CO₂ was not rescued by uracil or uridine supplementation while the auxotrophic phenotypes were reversed [11]. Therefore, Ura2 appears to buffer the cell against high levels of CO₂ by converting it into intermediates of the urea cycle. Together, these data indicate that host levels of CO₂ lead to substantial alteration of central carbon metabolism through multiple mechanisms.

Host concentrations of CO₂ alter C. neoformans cellular redox balance

In S. cerevisiae, acute depletion of glucose leads to an increase in mitochondrial respiration. Banerjee and colleagues have also demonstrated that glucose depletion in C. neoformans leads to increased oxidative stress which is a byproduct of mitochondrial activity [36]. Based on these precedents as well as the large set of mitochondrial genes required for CO₂ fitness, we predicted that host CO₂ concentrations would increase mitochondrial respiration and membrane potential. To test this hypothesis, we characterized the membrane potential of mitochondria in ambient and 5% CO₂ using JC-1 staining as previously reported by the Kronstad lab [37]. In this assay, increased mitochondrial membrane potential leads to a red shift in the JC-1 dye. As shown in Fig 6A, growth in 5% CO₂ leads to an increase in the proportion of red-shifted cells relative to ambient CO₂. This increase in mitochondrial membrane potential is consistent with an increase in respiration at host concentrations of CO₂ [38]. The membrane potential is driven by the TCA cycle and electron transport chain enzymes. Accordingly, deletion mutants of the TCA cycle enzyme succinyl CoA synthetase (LSC1) as well as three complex I/ubiquinone NADH dehydrogenase components CNAG_05267, CNAG_02403, and CNAG_02833 have reduced fitness in host CO₂ concentrations (Table 1). PFK1 and FBP26 reduce the ability of yeast to transition from glycolysis to respiration [39]. We, therefore, suggest this function may contribute to the reduced fitness of the pfk1∆ and fbp26∆ mutants (Table 1) at 5% CO₂.

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Fig 6. Host concentrations of CO₂ alter C. neoformans cellular redox balance and engage multiple acetyl-CoA generating pathways for tolerance.

A. The mitochondrial membrane potential of H99 in ambient or 5% CO₂ conditions for 24 h was assessed by JC-1 staining. JC-1 staining was analyzed by dividing the MFI resulting from excitation with the blue (488 nm) laser in the BL2 emission filter (574/26) by the MFI in the BL1 emission filter (530/30), i.e., red/green ratio of single yeast cells. Bars represent the average red/green ratio of a minimum of 10,000 cells per condition and error bars represent the SEM. Asterisks indicate a significant difference relative to H99 as determined by Student t test (P < 0.05). Cellular stores of NAD/NADH (B) and NADP+/NADPH (C) were determined for H99 after incubation in buffered RPMI medium at ambient or 5% CO₂ conditions for 24 h. Data are presented as ratios of the oxidized or reduced forms. Bars represent the average of three biological replicates and error bars represent SD. Asterisks indicate a significant difference relative to ambient conditions as determined by an unpaired t test (P < 0.05). D. The metabolic profile of H99 grown for 24 h in ambient air or 5% CO₂ was analyzed by tandem mass spectrometry. Data were processed and are presented for 6-P-Gluconate as normalized area. Asterisk indicates P < 0.05 for the comparison of ambient to 5% CO₂ using Welch corrected, two-tailed t test. E. Competitive fitness was determined for indicated strains after combining a 1:1 ratio of mNeonGreen-labeled H99 and unlabeled mutant cells in RPMI 1,640 medium with 165 mM MOPS at pH 7 and incubating at 37 °C in ambient air or 5% CO₂ for 24 h. Cell populations were characterized by flow cytometry, and the percentage of mNeonGreen-negative cells in 5% CO₂ was divided by those in ambient conditions to calculate competitive fitness. Bars represent the average of three biological replicates with error bars indicating SD. Asterisks indicate a significant difference relative to H99 as determined by one-way ANOVA with Tukey’s multiple comparisons test (P < 0.05). The competitive fitness of H99 and indicated strains (acl1∆ or yhm2∆) was determined in vivo after two inoculation routes in CD-1 mice. F. For intranasal inoculation (IN), wild-type and mutant cells were mixed in a 1:1 ratio prior to inoculation with 5 × 10⁴ CFU/animal in a 50 µL volume. Organs were harvested at day 10 post-inoculation. G. For intravenous inoculation (IV), wild-type and mutant cells were mixed in a 1:1 ratio prior to inoculation with 1 × 10⁵ CFU/animal in 100 µL volume. Organs were harvested at day 4 post-inoculation. Tissues were homogenized and serially diluted on YNB or YNB with 400 µg/mL hygromycin and incubated at 30 °C for two days before counting CFUs. The competitive fitness was determined by dividing the CFU of mutant cells (HYG resistant) by the CFU of H99 cells in lung (F) from the IN model or brain (G) from the IV model. Data are represented with the average CFU/mouse and SD. Asterisks indicate a significant difference relative to H99 as determined by unpaired t test (P < 0.05). The data underlying this Figure can be found in S3 Table and S1 Data.

https://doi.org/10.1371/journal.pbio.3003561.g006

To further characterize the effect of CO₂ on the redox state of C. neoformans, we measured the ratios of NAD+/NADH and NADP+/NADPH under ambient and host CO₂ concentrations. As shown in Fig 6B, CO₂ increases the NAD + /NADH ratio by ~2.5 fold. The increased NAD+/NADH ratio is consistent with the higher mitochondrial membrane potential observed in Fig 6A and further supports the conclusion that 5% CO₂ increases respiration and TCA cycle activity relative to ambient CO₂ concentrations. One of the consequences of increased respiration is increased oxidative stress in the cell. NADPH is a critical component of the biochemical processes that detoxify ROS [38]. Interestingly, the ratio of NADP+/NADPH is unchanged at 5% CO₂ compared to ambient CO₂ (Fig 6C). This suggests that CO₂-tolerant C. neoformans can compensate for the increase in respiration and oxidative state of the cell at 5% CO₂ without depleting NADPH stores significantly. The pentose-phosphate pathway generates two molecules of NADPH through the oxidation of glucose first to 6-phospho-gluconate and then, through an oxidative decarboxylation step, to ribulose-5-phosphate [40]. The levels of 6-phosphogluconate are reduced in 5% CO₂ relative to ambient CO₂ conditions which is consistent with increased pentose phosphate pathway activity (Fig 6D), possibly to maintain NADPH levels and compensate for increased ROS resulting from increased mitochondrial respiration.

Multiple acetyl-CoA-generating pathways are required for C. neoformans tolerance of host CO₂ concentrations

Acetyl-CoA is not only required for mitochondrial respiration through the TCA cycle but is also a fundamental substrate for lipid biosynthesis as well as for histone acetyltransferases involved in the regulation of gene expression [41]. Acetyl-CoA cannot diffuse across organelle membranes and, therefore, distinct pools of acetyl-CoA are present in the mitochondria, cytosol, and nucleus [41]. In the presence of glucose, glycolysis delivers pyruvate-derived acetyl-CoA to the mitochondria and TCA cycle. Our metabolomics data suggests that glucose and glycolysis are reduced at 5% CO₂ and that ketogenic amino acids are mobilized to provide acetyl-CoA to support the increased mitochondrial respiration driven by the acute reduction in glucose.

Lipid catabolism through β-oxidation contributes to the acetyl-CoA pool and, therefore, could also contribute to acetyl-CoA homeostasis during CO₂ stress [32]. Supporting this hypothesis, we identified three mutants in the β-oxidation pathway that have reduced CO₂ fitness: the enoyl CoA hydratase CNAG_04531, the multi-functional protein MFE2, and the 3-oxo-acyl CoA thiolase POT1 (Table 1). Enoyl-CoA hydratases catalyze the second step of fatty acid β-oxidation while Pot1 directly generates acetyl-CoA. MFE2 is a peroxisomal enzyme that catalyzes the first two steps of β-oxidation [38]. MFE2 is subject to glucose repression in C. neoformans, and POT1 has been shown to be negatively regulated by glucose in S. cerevisiae [42]. The expression of both MFE2 (1.8-fold, FDR = 9e-15) and POT1 (3-fold, FDR 1e-68) is increased in 5% CO₂, further supporting its likely role in generating acetyl-CoA under these conditions [12]. In addition, β-oxidation is positively regulated by the AMP-activated protein kinase Snf1 in other systems [44], and this kinase is required for full fitness at host CO₂ concentrations (competitive index: 0.67; p = 0.012, S1 Table). Finally, POT1, MFE2, and enoyl CoA hydratase CNAG_04531 are required for full fitness during infection [14]. Thus, the in vivo fitness defects of genes related to β-oxidation may be due, at least in part, to their importance in the adaptation of C. neoformans to host CO₂.

Two additional enzymes could also contribute to the generation of acetyl-CoA: acetyl-CoA synthetase (ACS1) and aceto-acetyl-CoA synthetase (KBC1, [45]). Acs1 converts acetate to acetyl-CoA. Kbc1 condenses the ketone body acetoacetate with CoA to give aceto-acetyl-CoA which, in turn, is a substrate for Pot1, leading to two molecules of acetyl-CoA. We, therefore, tested the CO₂ fitness of acs1∆, kbc1∆, and the corresponding double deletion mutants using the competition assay. The kbc1∆ acs1∆ double mutant showed a modest reduction in CO₂ fitness whereas neither single mutant differed from WT (Fig 6E), suggesting that these enzymes play a minor role in maintaining acetyl-CoA homeostasis during CO₂ stress.

Amino acid catabolism and β-oxidation to acetyl-CoA occur in the mitochondria and directly provide acetyl-CoA for use in the TCA cycle. However, acetyl-CoA is also required in other cellular components to support biosynthesis and gene expression. An important source of cytosolic and nuclear acetyl-CoA in C. neoformans and other cells is, in fact, the mitochondria. Indeed, previous work by the Kronstad lab showed that the majority of cytosolic and nuclear acetyl-CoA appears to be derived from citrate generated in the mitochondria [46]. Citrate is exported to the cytosol, where it is converted to acetyl-CoA and oxaloacetate by ATP-citrate lyase (ACL1). In prior work, we found that the acl1∆ mutant has a severe growth defect in 5% CO₂ [45]. This suggests that nuclear and cytosolic acetyl-CoA stores remain primarily dependent on Acl1.

The deletion mutant of a tricarboxylic acid transporter, CNAG_06096, has a strong fitness defect in 5% CO₂ (Table 1). CNAG_06096 is homologous to S. cerevisiae Yhm2 which is a mitochondrial citrate-oxoglutarate antiporter [47]. In other fungi with ATP-citrate lyase enzymes such as Aspergillus spp., deletion of the citrate-oxoglutarate transporter leads to reduced acetyl-CoA production [48]. Presumably, this is due to reduced cytosolic citrate in the absence of the transporter. If Yhm2 plays this role during mammalian C. neoformans infection, then, like the acl1∆ mutant, it should have a profound fitness defect in vivo. To test this hypothesis, we infected mice with a 1:1 mixture of H99 and either the acl1∆ or yhm2∆ mutant via inhalation (Fig 6F) and intravenous injection (Fig 6G). As expected, based on the previous report of its reduced virulence, the acl1∆ had dramatically reduced fitness in both lung and brain tissue. The yhm2∆ mutant showed a similarly profound reduction in fitness relative to WT. These data indicate that mitochondrial citrate-derived acetyl-CoA is required for C. neoformans tolerance of CO₂ and infection of mammals.

Taken together, these data indicate that host concentrations of CO₂ establish a relatively low glucose state in C. neoformans. In response, oxidation of fatty acids and amino acid catabolism generates mitochondrial acetyl-CoA which then supports increased respiratory metabolism. Cytosolic and nuclear pools of acetyl-CoA remain largely dependent on the export of mitochondrial citrate through the transporter Yhm2; citrate is then converted to acetyl-CoA by Acl1 in the nucleus and cytosol.

The glutathione and TOR-Ypk1 pathways buffer C. neoformans against CO₂-induced ROS accumulation

One of the consequences of increased mitochondrial respiration is the generation of ROS. Glutathione is an important mechanism by which ROS species are detoxified. Recently, Black and colleagues described the importance of glutathione-mediated redox regulation in the virulence of C. neoformans through effects on melanization and titan cell formation [49]. In addition, loss of glutathione affects a variety of metabolic pathways and reduces fitness in low-nutrient growth conditions. The glutathione synthetases GSH1 and GSH2 are both required for CO₂ tolerance in H99 (Fig 7A). The biosynthesis of glutathione is integrated with sulfate assimilation as well as methionine and cysteine biosynthesis (Fig 7B, [50]). Consistent with this integration, five genes in the sulfur assimilation pathway (SUL1, MET3, MET5, MET10) are required for CO₂ tolerance along with CYS4 which converts homocysteine to cystathionine and MET2 which generates the homoserine that is coupled with cystathionine to generate cysteine (Fig 7A and 7B). Homoserine levels are significantly reduced under 5% CO₂ (Fig 7C), further supporting the notion that this pathway is highly active under those conditions.

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Fig 7. The glutathione and TOR-Ypk1 pathways buffer C. neoformans against CO₂-induced ROS accumulation.

A. Competitive fitness was determined for indicated strains after combining a 1:1 ratio of mNeonGreen-labeled H99 and unlabeled mutant cells in RPMI 1,640 medium with 165 mM MOPS at pH 7 and incubating at 37 °C in ambient air or 5% CO₂ for 24 h. Cell populations were characterized by flow cytometry, and the percentage of mNeonGreen-negative cells in 5% CO₂ was divided by those in ambient conditions to calculate competitive fitness. Bars represent the average of three biological replicates with error bars indicating SD. Asterisks indicate a significant difference relative to H99 as determined by Student t test (P < 0.05). B. Schematic of the glutathione biosynthesis pathway in C. neoformans. Genes required for CO₂ fitness are indicated in red. Created in BioRender.com. Ristow, L. (2026) https://BioRender.com/f5i198sc.C. The metabolic profile of H99 grown for 24 h in ambient air or 5% CO₂was analyzed by tandem mass spectrometry. Data were processed and are presented for homoserine as normalized area. The normalized area of each metabolite was determined by tandem mass spectrometry as described in materials and methods for H99 incubated in buffered RPMI in ambient air or 5% CO₂. Asterisks indicate statistically significant differences in abundance at 5% CO₂ relative to ambient air (Welch corrected, two-tailed t test, P < 0.05). D. Competitive fitness of the glutathione biosynthesis pathway mutants was determined as in (A) with the following modification: cells were grown in RPMI 1,640 medium with 165 mM MOPS at pH 7 alone or with the addition of 200 µM cysteine or 1 mM glutathione for 24 h in 5% CO₂. The percentage of mNeonGreen negative cells in media with supplement was divided by the percentage of mNeonGreen negative cells without supplement to determine competitive fitness. Bars indicate the average of three biological replicates with error bars indicating SD. All conditions tested were significantly different relative to unsupplemented cells for each mutant as determined by Student t test (P < 0.05). E&F. ROS activity of indicated strains after growth in ambient air or 5% CO₂ for 24 h was assessed by DCFDA fluorescence assay. The average MFI of three biological replicates for each strain and condition is represented with error bars indicating SD. Asterisks indicate a significant difference as determined by one-way ANOVA with Sidak’s multiple comparisons test (P < 0.05). The data underlying this Figure can be found in S1 and S3 Tables and S1 Data.

https://doi.org/10.1371/journal.pbio.3003561.g007

To explore the role of these pathways in CO₂ tolerance, we compared the CO₂ fitness of each mutant in the presence and absence of supplemental cysteine and glutathione (Fig 7D). The CO₂ fitness of each mutant was improved in the presence of both cysteine and glutathione. Black and colleagues had found that reduced glutathione had effects on the cell beyond its role in detoxifying ROS [49]. We, therefore, used the DCFDA assay to characterize the levels of ROS induced by CO₂ in the WT strain and the gsh2∆ mutant (Fig 6E). In WT cells, 5% CO₂ does not induce a significant increase in ROS. However, cellular ROS increases significantly in the gsh2∆ mutant relative to WT in 5% CO₂ (Fig 7E). These data suggest that host concentrations of CO₂ cause oxidative stress in C. neoformans but that CO₂-tolerant strains such as H99 utilize glutathione to detoxify these species as part of a compensatory response. It is also possible that glutathione is required to maintain metabolic homeostasis as well. Taken together, these data indicate that the reduced in vivo fitness of genes involved in glutathione homeostasis may also be due to their role in the response to increased CO₂ concentrations encountered by C. neoformans during mammalian infection.

In S. cerevisiae, the TOR-Ypk1 pathway also buffers the cell from ROS generated in the mitochondria and vacuole through its regulation of sphingolipid biosynthesis [51]. In C. neoformans, TOR-Ypk1 is required for sphingolipid biosynthesis, and we hypothesized that this pathway may also buffer the cell against CO₂-induced ROS. High levels of ROS in ambient CO₂ are present in the ypk1∆ mutant relative to WT (Fig 7F). This observation is consistent with the model proposed by Niles and colleagues in which TOR-Ypk1 suppresses the accumulation of ROS in S. cerevisiae [51]. The levels of ROS in the ypk1∆ mutant do not increase further in 5% CO₂. However, the loss of ROS homeostasis in the ypk1∆ mutant seems likely to contribute to its susceptibility to 5% CO₂. As discussed above Ypk1 is required for synthesis of sphingolipids in C. neoformans and other fungi through regulation of SPT [22,23]. Niles and colleagues also found that reduced sphingolipid biosynthesis increased cellular ROS [51]. If this mechanism is also operative at host levels of CO₂ in C. neoformans, then the gcs1∆ mutant would be expected to have the same phenotype. Although loss of GCS1 does not alter ROS levels in ambient CO₂ concentrations (Fig 7F), ROS levels are increased in the gcs1∆ mutant at 5% CO₂. Thus, the TOR-Ypk1 pathway is required for the prevention of toxic levels of cellular ROS, and its regulation of sphingolipid biosynthesis is likely to contribute to this function. Furthermore, the glucosylceramide arm of the sphingolipid pathway directly contributes to the buffering of ROS generated by C. neoformans growth at host concentrations of CO₂.

Ethics statement

This study was performed according to the guidelines of the National Institutes of Health and the University of Georgia Institutional Animal Care and Use Committee (IACUC). The animal models and procedures used have been approved by the IACUC (AUP protocol number: A2023 03-033-A3).

Strains and growth conditions

Yeast extract-peptone-2% dextrose (YPD) was prepared according to standard recipes [56]. RPMI 1,640 without glutamine or sodium bicarbonate was buffered with 165 mM MOPS and pH adjusted to 7. Strains used in this work are described in S4 Table. The C. neoformans deletion library (2015, 2016, 2020 Madhani plates) was acquired from the Fungal Genetic Stock Center (FGSC). Strains were stored at −80 °C in 20% glycerol. Frozen stocks were recovered on solid YPD medium at 30 °C for 2 days. To prepare for assays, 3 mL YPD was inoculated per strain and grown overnight, 30 °C, shaking at 200 rpm.

Strain construction

For generation of an independent yhm2∆ strain (S4 Table), CRISPR/Cas9 short-arm homology (SAH) and transient CRISPR-Cas9 coupled with electroporation (TRACE) methods were used as published [57,58]. CRISPR components were PCR purified according to manufacturer instructions (QIAquick PCR Purification Kit; cat no. 28104) and transformed into H99 via electroporation using the “Pic” setting on a Bio-Rad Micropulser. For gene deletion constructs, YHM2 SAH Repair 5′ and 3′ oligos designed with a 50 bp sequence matching the flanks of the YHM2 coding sequence and a 20 bp sequence matching the flanks of the hygromycin (HygB) resistance marker cassette were used to amplify a repair construct, which replaced the entire open reading frame for the target gene with the HYGB cassette. Gene deletion constructs were transformed along with Cas9 DNA and two gene-specific sgRNAs into H99. Transformants were selected on YPD medium with 400 µg/mL hygromycin B, and knockouts were PCR verified at the 5′ flank with LCR031 and gene-specific “5′ KO confirmation primer”, at the 3′ flank with LCR323 and gene-specific “3′ KO confirmation primer”, and for lack of the native locus with gene-specific “orf confirmation” primer pairs (S5 Table).

Flow cytometry-based competitive growth assay

The Madhani mutant collection was screened using a competitive growth assay in which each mutant was compared to the H99 reference strain following previously published procedures [12]. Briefly, 96-well plates were prepared with 200 µL YPD per well. Wells were inoculated from growth on solid YPD plates with individual strains to be tested and the reference strain H99-mNeonGreen and grown statically overnight at 30 °C. Dilutions were performed to standardize input of unlabeled strains and H99-mNeonGreen in a 1:1 ratio, at 2 × 10⁴ cells/mL final concentration in RPMI 1,640 medium with 165 mM MOPS, pH 7. Co-cultures were grown at 37 °C for 24 h in ambient air or 5% CO₂ before analyzing on an Attune NxT Flow Cytometer with CytKick autosampler and Attune Cytometric software. mNeonGreen positive and negative populations were identified by histogram plot with 10,000 single cells counted per sample. The percent of mNeonGreen negative cells in CO₂ conditions was divided by the percent of mNeonGreen negative cells in ambient conditions to determine the competitive fitness in CO₂ relative to ambient conditions. Statistical significance was determined by chi-squared test in Microsoft Excel. In experiments to follow up the screen, the following additions to the medium were made as indicated at the following final concentrations; cysteine (200 µM), glutathione (1 mM), and the final analysis compared mutant cells with supplementation to those without supplementation at 37 °C, 5% CO₂.

Solid agar growth assays

Cells from overnight cultures of indicated strains were washed twice with PBS prior to quantifying OD₆₀₀. Strains were diluted to an OD₆₀₀:1, followed by 10-fold serial dilutions. Three µL from each dilution was spotted on agar plates and grown inverted at 37 °C in ambient air or at 10% CO₂ on YPD plates or YPD buffered with HEPES at pH 8. Images were captured at 48 h.

Aureobasidin A susceptibility assay

Cells from overnight cultures were washed twice with PBS prior to quantifying concentration on a Countess II FL automated cell counter. Cells were diluted to 1 × 10⁷ cells/mL to plate on RPMI 1,640 medium with 165 mM MOPS, pH 7 agar plates. A sterile q-tip was saturated in the cell solution before streaking an entire plate to generate a lawn. When plates had dried, a sterile filter paper dot was placed in the center of the plate. Aureobasidin A was added in a 20 µL volume at a final concentration of 100 µg/disk. Plates were incubated at 37 °C in ambient air or 5% CO₂. Images were captured at 48 h.

Metabolomic analysis

A 30 mL culture of YPD was inoculated with H99 and grown overnight at 30 °C in a 220 rpm shaking incubator. Simultaneously, two groups of 4 flasks containing 50 mL of RPMI media buffered to pH 7 with 165 mM MOPS were pre-equilibrated by shaking at 220 rpm in a 37 °C incubator with either ambient air or 5% CO₂ overnight. The following day, late log-phase cultures were back-diluted to an OD₆₀₀ of 0.5 and allowed to double at 37 °C in a 220 rpm shaking incubator. Pre-equilibrated flasks were inoculated to an initial OD₆₀₀ of 0.1 from the log phase culture and allowed to grow for 24 h. 100 ODs were collected from each flask, cell pellet rinsed with ice-cold UltraPure Distilled Water (Invitrogen), then flash frozen in an ethanol dry-ice bath and stored at −80 °C. For extraction, cell pellets were subjected to three rounds of homogenization with glass beads for one min. Homogenates were resuspended in 2 mL of methanol and transferred to 8 mL glass vials containing 4 mL of cold chloroform (LiChrosolv, Sigma). Samples were then vortexed for 1 min and 2 mL of water (Omnisolv, Sigma) was added. Samples were vortexed for 1 min and then subjected to 10 min of centrifugation at 3,000g for phase separation. The aqueous phase was collected (~3.8 mL) and transferred to a new glass vial. Samples were frozen at −80 °C and submitted to the Harvard Center for Mass Spectrometry (HCMS) where they were dried under nitrogen flow and resuspended in 100 µL of 30% acetonitrile in water.

Untargeted metabolomics was performed using HILIC-UHPLC-MS/MS. A 5 µl injection was separated on a HILICON iHILIC-P Classic column (150 × 2.1 mm, 5 µm) at 40 °C with a ThermoFisher IDX system with HESI source in positive and negative polarity (resolution 120,000; AGC target 1 × 105; m/z 65–1,000). The 45 min LC gradient (0.15 mL/min after 0.5 min ramp from 0.05 mL/min) used mobile phase A (20 mM ammonium carbonate, 0.1% ammonium hydroxide in water) and B (97% acetonitrile in water): 93% B (0–0.5 min), linear to 40% B (19 min), to 0% B (28–33 min), return to 93% B (36–45 min). Eluent was diverted to waste at 0–0.5 min and 32–46 min. A pooled QC sample underwent AcquireX deepscan (5 levels) in both polarities for MS/MS.

Data were processed in Compound Discoverer 3.2 with the following workflow: (1) MS1 peak detection and integration (threshold-based); (2) retention time alignment; (3) gap-filling by re-integration of low-intensity or noise regions; (4) adduct grouping into single features (neutral monoisotopic mass reported); (5) background subtraction using blanks; (6) median-centered normalization; (7) formula prediction from accurate mass, isotopic pattern, and MS2 (if available); (8) identification via mzVault (local, RT-inclusive) then mzCloud (online MS2); and (9) manual curation of IDs and integrations. One ambient sample exhibited anomalous total intensity and was excluded from differential analysis; normalized areas without this anomalous sample are reported. Peak areas were median-centered and log-transformed prior to statistical analysis, where a Welch corrected, two-tailed t test was performed to compare Ambient (n = 3) versus Elevated (n = 4) conditions. Features with log2 (fold-change) >1 and P < 0.05 were considered significantly regulated.

Mitochondrial membrane potential and reactive oxygen species assays

Cells from overnight cultures of indicated strains were washed twice with PBS prior to quantifying concentration on a Countess II FL automated cell counter. Cells were diluted to 7.5 × 10⁵ cells/mL in RPMI 1,640 medium with 165 mM MOPS, pH 7 and aliquoted in 3 ml volumes in 6-well plates, with 3 wells per replicate, performed in biological triplicate. Plates were incubated at 37 °C in ambient air or 5% CO₂ for 24 h. At 24 h, replicate wells were combined, concentration was quantified by cell counter, and 1 × 10⁶ cells were aliquoted into a 24-well plate per staining condition. Cells were resuspended in 1 ml RPMI containing 2.5 µM JC-1 (catalog no. M34152; Thermo Fisher) or 16 µM DCFDA (catalog no. D6883; Sigma) and incubated at 37 °C in ambient air or 5% CO₂ for 1 h. Cells were washed once with PBS, then resuspended in 500 µl PBS and analyzed on an Attune NxT Flow Cytometer with Attune Cytometric software. JC-1 staining was analyzed by dividing the mean fluorescence intensity (MFI) resulting from excitation with the blue (488 nm) laser in the BL2 emission filter (574/26) by the MFI in the BL1 emission filter (530/30), i.e., red/green ratio of single yeast cells. DCFDA staining was analyzed by measuring the MFI of single yeast cells in the BL1 emission filter (530/30) following excitation with the blue laser.

Quantification of NAD/NADH and NADP+/NADPH ratios

Cells from overnight cultures of H99 were washed twice with PBS prior to quantifying concentration on a Countess II FL automated cell counter. Cells were diluted to 7.5 × 10⁵ cells/mL in RPMI 1,640 medium with 165 mM MOPS, pH 7 and aliquoted in 3 ml volumes in 6-well plates, with 3 wells per replicate, performed in biological triplicate. Plates were incubated at 37 °C in ambient air or 5% CO₂ for 24 h. At 24 h, replicate wells were combined, concentration was quantified by cell counter, and two tubes of 1 × 10⁷ cells were pelleted per replicate/condition to allow simultaneous analysis of the oxidized and reduced form from each replicate. Cells were frozen at −80 °C and lyophilized overnight. Manufacturer kit instructions were followed for NAD+/NADH (BioAssay E2ND-100) or NADP + /NADPH (BioAssay Systems E2NP-100) quantification.

Mouse model of cryptococcosis

CD1 female mice (022CD1) with a body weight of 18–25 g were purchased from Charles River. The mice were housed with a max of 5 per cage with a 7am–7 pm 12 h dark/12 h light cycle. They were fed LabDiet 5053 irradiated Picolab Rodent Diet 20. The water was provided by the automated watering system connected to the rack, which has several filtration systems. The ambient temperature was between 20 and 24 °C with 30%–70% humidity. For intravenous inoculation (IV), wild-type and mutant cells were mixed in a 1:1 ratio prior to inoculation with 1 × 1 0⁵ colony-forming units (CFUs)/animal in 100 µl volume. Organs were harvested at day 4 and homogenized in 2 mL of cold sterile PBS using an IKA-T18 homogenizer. Homogenized organs were serially diluted, plated onto YNB and YNB with 400 µg/mL hygromycin, and incubated at 30 °C for 2 days before counting CFUs. Neither the acl1Δ nor yhm2Δ strains were detected in the brain. For intranasal inoculation (IN), wild-type and mutant cells were mixed in a 1:1 ratio prior to inoculation with 5 × 10⁴ CFU/animal in a 50 µl volume. Organs were harvested at day 10, and fungal burden quantification was performed as above.

Western blotting

Overnight cultures of H99 were rinsed in water and resuspended to spot 5 × 10⁷ cells in 20 µl in biological triplicate on an RPMI 1,640 medium with 165 mM MOPS, pH 7 agar plate. Replicate plates were incubated at 37 °C overnight. Prior to CO₂ exposure, a 0 h time point was collected, then one plate was shifted to 37 °C + 5% CO₂, while one remained at 37 °C ambient CO₂ for 2 h. At 2 h, all replicates were swabbed from the plates into individual tubes containing 500 µl 0.2 N NaOH. Suspensions were incubated at room temperature for five minutes. Cells were pelleted and resuspended in 20 µl reducing sample buffer. Samples were boiled for 5 min, then pelleted at 5,000 rpm for 3 min before removing supernatant to fresh tubes. Ten µl from each replicate was loaded into duplicate 10% SDS-PAGE gels and run at 80 V. Samples were transferred to a nitrocellulose membrane for 1 h at 100 V. Membranes were blocked with 5% bovine serum albumin (BSA) in tris-buffered saline with 0.1% tween 20 (TBST) for 1 h at RT, then incubated with 1:10,000 rabbit anti-Hog1 (generous gift from Yong-Sun Bahn) or rabbit anti-phospho-p38 MAPK (catalog no. 4092S; Cell Signaling Technology) at 1:10,000 in 5% BSA/TBST overnight at 4 °C. Membranes were washed 3 times for 5 min with TBST, then incubated for 1 h at RT with 1:10,000 goat anti-rabbit horseradish peroxidase (HRP) (catalog no. STAR208P; Bio-Rad) in 5% BSA/TBST. Membranes were washed 3 times for 5 min with TBST, then developed with chemiluminescent substrate (catalog no. 1705060; BioRad) and imaged on a myECL imager (ThermoFisher).

Statistical analyses

Data were graphed and analyzed in GraphPad Prism 9 software or Microsoft Excel. Statistical analysis and graph descriptions, including ` values are provided in figure legends.