Abstract
The evolution of pathogens in response to selective pressures present during chronic infections can influence persistence, virulence, and the outcomes of antimicrobial therapy. Because subpopulations within an infection can be spatially separated and the host environment can fluctuate, an appreciation of the pathways under selection may be most easily revealed through the analysis of numerous isolates from single infections. Here, we continued our analysis of a set of clonally-derived Clavispora (Candida) lusitaniae isolates from a single chronic lung infection with a striking enrichment in the number of alleles of MRR1. Genetic and genomic analyses found evidence for repeated acquisition of gain-of-function mutations that conferred constitutive Mrr1 activity. In the same population, there were multiple alleles with both gain-of-function mutations and secondary suppressor mutations that either attenuated or abolished the constitutive activity suggesting the presence of counteracting selective pressures. Our studies demonstrated tradeoffs between high Mrr1 activity, which confers resistance to the antifungal fluconazole, host factors, and bacterial products through its regulation of MDR1, and resistance to hydrogen peroxide, a reactive oxygen species produced in the neutrophilic environment associated with this infection. This inverse correlation between high Mrr1 activity and hydrogen peroxide resistance was observed in multiple Candida species and in serial analysis of populations from this individual collected over three years. These data lead us to propose that dynamic or variable selective pressures can be reflected in population genomics and that these dynamics can complicate the drug resistance profile of the population.
Importance Understanding microbial evolution within patients is critical for managing chronic infections and understanding host-pathogen interactions. Here, our analysis of multiple MRR1 alleles in isolates from a single Clavispora (Candida) lusitaniae infection revealed the selection for both high and low Mrr1 activity. Our studies reveal tradeoffs between high Mrr1 activity, which confers resistance to the commonly used antifungal fluconazole, host antimicrobial peptides and bacterial products, and resistance to hydrogen peroxide. This work suggests that spatial or temporal differences within chronic infections can support a large amount of dynamic and parallel evolution, and that Mrr1 activity is under both positive and negative selective pressure to balance different traits that are important for microbial survival.
Introduction
Understanding the positive and negative selective pressures that shape drug resistance profiles in microbial populations is critical for combating the development of antimicrobial resistance, an ever-increasing problem in clinical settings. Increased drug resistance in bacteria and fungi has been associated with clinically- and agriculturally-used antimicrobial agents (reviewed in (1–3)), and drug resistance elements may be selected for based on their ability to protect against factors produced by other microbes or plant, animal, and insect hosts (4, 5). Based on the analysis of bacterial isolates of Burkholderia dolosa or Pseudomonas aeruginosa from single patients and across cohorts of patients, it is clear that in vivo factors can lead to the repeated selection for subpopulations with the same genes or pathways mutated (6–8). Furthermore, there is evidence that pathways can be upregulated then downregulated in the same phylogenetic lineages. For example, suppressor mutations within P. aeruginosa algU frequently arise in strains with high AlgU signaling caused by mutations in the gene encoding the AlgU repressor MucA (9). Less is known about the negative selective pressures acting against sustained microbial resistance.
In Demers et al. (10), we described a set of twenty recently-diverged Clavispora (Candida) lusitaniae isolates obtained from the lung infection of a single individual with cystic fibrosis (CF). C. lusitaniae is among the emerging non-albicans Candida spp. that cause life threatening disseminated infections in individuals who are immunocompromised (11, 12), and can cause infections of the gastrointestinal tract (13–15), surgical sites, or implanted devices in immunocompetent individuals. C. lusitaniae is notorious for its rapid development of resistance to antifungal drugs including amphotericin B, azoles and echinocandins (14, 16–19) and, relative to Candida albicans and other Candida species that are both opportunistic pathogens and members of the mycobiome, it is more closely related to Candida auris, a species in which multi-drug resistant isolates have caused hospital-associated outbreaks (20–26). Our previous analyses of heterogeneity in fluconazole (FLZ) resistance among these isolates identified numerous distinct alleles of MRR1 (CLUG_00542). Multiple alleles encoded gain-of-function (GOF) mutations causing constitutive Mrr1 activity, which, as in other Candida species, increased expression of MDR1 and Mdr1 multidrug efflux pump activity (10, 27–32). At the time that these isolates were recovered, the patient had no history of antifungal treatment, suggesting that selection for constitutively active Mrr1 variants may have been driven by the need for resistance to other host- or microbe-produced compounds. Within this study, however, we found multiple lineages with recently evolved MRR1 alleles that rendered cells more sensitive to FLZ than even mrr1Δ strains. Here, we address the perplexing question of why this population had recently diverged MRR1 alleles that encoded both high and low Mrr1 activity. To do so, we expressed both native and synthesized MRR1 alleles that represent intermediates during MRR1 evolution in a common genetic background and tested the effects of these alleles on growth in in vivo relevant conditions. Using genetics and genomics, we concluded that multiple C. lusitaniae MRR1 alleles that conferred low Mrr1 activity resulted from initial mutation that caused constitutive Mrr1 activity followed by a second mutation that either suppressed constitutive activation or inactivated the protein. Constitutive Mrr1 activity caused increased sensitivity to a variety of biologically relevant compounds including hydrogen peroxide (H2O2) and suppression of constitutive Mrr1 activity rescued growth under many of these conditions. Monitoring populations from respiratory samples from this subject over time supports the model that there are opposing selective pressures in vivo that select for and against constitutive Mrr1 activity, as reflected by the tradeoff between FLZ and H2O2 resistance seen over time. These data explain the persistence of a heterogeneous fungal population and underscores the complexity and parallelism of evolution that is possible in the human lung during chronic disease.
Results
Naturally evolved C. lusitaniae MRR1 alleles confer altered Mrr1 activity and FLZ resistance
Each of the twenty closely-related C. lusitaniae isolates from a single individual contained at least one nonsynonymous single nucleotide polymorphism (SNP) or single nucleotide insertion or deletion (indel) in MRR1 relative to the deduced MRR1 sequence of their most recent common ancestor (MRR1ancestral) (Fig. 1A) (10). To determine the impact of specific mutations in MRR1 on Mrr1 activity, we expressed different MRR1 alleles in a common genetic background in which the native MRR1 had been deleted (U04 mrr1Δ). Deletion of MRR1 in the FLZ-resistant strain U04 reduced the FLZ minimum inhibitory concentration (MIC) from 32 μg/ml to 4 μg/ml (10) and the decrease in MIC was complemented by restoring the native MRR1Y813C allele (Fig. 1B). Complementation of U04 mrr1Δ with the MRR1ancestral allele led to a FLZ MIC of 1 μg/ml which was 4-fold lower (P<0.0001) than the FLZ MIC of U04 mrr1Δ, suggesting that MRR1ancestral had a function that reduced the FLZ MIC (Fig. 1B). Expression of an MRR1 allele from a FLZ-sensitive isolate in the population (MRR1L1191H+Q1197*) also reduced the FLZ MIC to levels comparable to those for MRR1ancestral (0.5-1 μg/ml) (Fig. 1B). Similar correlations between MRR1 allele and FLZ MIC were observed when the MRR1ancestral, MRR1Y813C, and MRR1L1191H+Q1197* alleles were expressed in a mrr1Δ derivative of the FLZ-sensitive strain U05, which indicated that strain background did not contribute to the FLZ MIC conferred by different MRR1 alleles (Fig. 1C). We previously published that FLZ resistance correlated with expression of MDR1 (10), also referred to as MFS7 (19). Deletion of MDR1 reduced the MIC, and the MIC was even lower in U04 mrr1Δ mdr1Δ (Fig. S1A) indicating that the moderately higher levels of FLZ resistance in U04 mrr1Δ compared to a strain with MRR1ancestral was MDR1-dependent.
RNA-sequencing (RNA-seq) analysis validated the previously published result that MDR1 correlated with the FLZ MIC for the different Mrr1 variants (10). The expression of MGD1 and MGD2, two C. lusitaniae genes shown to be Mrr1-regulated, correlated with the expression of MDR1 (Fig. 1D and Table S1) (10, 13, 33). Gene expression differences between U04 (MRR1Y813C), U04 mrr1Δ, and U04 mrr1Δ +MRR1Y813C found that mrr1Δ is fully complemented upon return of MRR1Y813C to the native locus (Fig. 1D and Fig. S2A for correlation plot) and that Mrr1 positively and negative regulates a large set of genes. Furthermore, a correlation analysis found that U04 mrr1Δ +MRR1ancestral and U04 mrr1Δ +MRR1L1191H+Q1197* were similar to each other but distinct from the mrr1Δ (Fig. S2A and 1B). A linear model comparing these strains identified forty-one genes with at least a 2-fold change in expression and corrected P value <0.05 (FDR). Comparison of non-isogenic C. lusitaniae strains similarly identified at least fourteen of the genes in Table S1 as putatively Mrr1-regulated (10, 13). Eighteen genes were homologs or had similar predicted functions as genes previously published as regulated by C. albicans Mrr1 (29), including MDR1, FLU1 and multiple putative methylglyoxal reductases encoded by GRP2-like genes, such as MGD1 and MGD2 (Fig. 1D and Table S1). Other genes within the Mrr1 regulon are discussed further below.
The unexpected finding that FLZ MIC was higher upon deletion of MRR1 relative to a strain with MRR1ancestral or an allele from a FLZ-sensitive strain was also observed in distantly-related C. lusitaniae strains, ATCC 42720 and DH2383 (FLZ MICs of ~1-2 μg/ml). In both cases, deletion of MRR1 led to a 2-4-fold increase in FLZ MIC to 4-8 μg/ml (Fig. S1B, P<0.001). The increase in FLZ MIC in mrr1Δ strains was not due to introduction of the selectable marker, NAT1, which encodes a nourseothricin acetyltransferase (34), as expression of NAT1 from an intergenic site in the FLZ-sensitive U05 strain did not increase the FLZ MIC (Fig. S1C). These data led us to hypothesize that some Mrr1 variants (MRR1Y813C) lead to high Mdr1 activity while other Mrr1 variants (both MRR1ancestral and the recently-diverged MRR1L1191H+Q1197* alleles) repressed the expression of at least some Mrr1-controlled genes, such as MDR1. Indeed, the RNA-Seq analysis identified six genes, including MDR1, that while positively regulated when Mrr1 was constitutively active, were more highly expressed in U04 mrr1Δ than those strains encoding low activity Mrr1 variants (Fig. 1D and S2B). These data suggest that, for a small subset of Mrr1-regulated genes, including MDR1, low activity Mrr1 variants directly or indirectly inhibit gene expression.
Truncation of MRR1 has varied effects on Mrr1 activity and inducibility in clinical isolates
All twenty sequenced clinical C. lusitaniae isolates from a single human subject (Fig. 1A) had MRR1 alleles with either one or two nonsynonymous mutations relative to MRR1ancestral, and we found that C. lusitaniae isolates with two mutations in MRR1 had a significantly lower average FLZ MIC than isolates with a single MRR1 mutation (Fig. 2A, P<0.001) (10). Interestingly, six of the seven MRR1 alleles in the “two mutation” set encoded premature stop codons, resulting in loss of 34-906 amino acids (Fig. 2B). There were two instances in which the same mutation was found with different nonsense mutations (*) or single nucleotide indels that led to early termination (t): MRR1Y1126N+P1174P(t) or MRR1Y1126N+S359*, and MRR1R1066S+K912N(t), MRR1R1066S+Y1061* or MRR1R1066S+G1231* (common mutation in bold, Fig. 1A) suggesting a complex evolutionary history for these alleles.
To better understand the effects of MRR1 mutations on Mrr1 activity, we analyzed the effects of a chemical inducer of Mrr1 activity, benomyl (35–37), on MDR1 expression. Benomyl strongly induced MDR1 expression in an Mrr1-dependent manner in the FLZ-sensitive strain ATCC 42720 (Fig. 2C) and, to a lesser extent, in the FLZ-resistant strain U04, which has high basal MDR1 expression (Fig. 2D) (10). Quantitative RT-PCR analysis of MDR1 expression and induction by benomyl in this collection of clinical isolates with different Mrr1 variants found that the two isolates with the lowest basal MDR1 expression and lowest FLZ MIC (U05 and U07) had the greatest induction by benomyl (34- and 27-fold, respectively) (Fig. 2D). Three isolates, L11, L12 and U06, had intermediate FLZ MICs and MDR1 expression levels, and did not show benomyl induction, similar to mrr1Δ, and all encoded Mrr1 variants lacking greater than 200 amino acids leading us to propose that these mutations caused a loss of Mrr1 function (Fig. 2D). Other isolates showed a correlation between higher basal MDR1 levels and elevated FLZ MICs, and this pattern was associated with lower relative levels of benomyl induction (Fig. 2D).
Premature stop codons repeatedly arose in constitutively active Mrr1 variants and caused either a loss of constitutive Mrr1 activity or complete loss of function
In light of the mixed effects that these two-mutation MRR1 alleles had on Mrr1 activity, we sought to determine the individual effects of mutations within each allele with a focus on the two strains with the lowest basal MDR1 expression and the strongest induction of MDR1 in response to benomyl: MRR1L1191H+Q1197* (in U05) and MRR1Y1126N+P1174P(t) (in U07) (Fig. 3A and 3B). We found that the MRR1L1191H mutation caused a 32-fold increase in FLZ MIC (Fig. 3C) and 22-fold increase in MDR1 expression (Fig. 3D) compared to MRR1ancestral indicating that, like the Mrr1-Y813C variant, Mrr1-L1191H was constitutively active. In contrast, MRR1Q1197*, which caused the loss of 68 amino acids from the C-terminus of Mrr1, did not significantly alter the FLZ MIC compared to MRR1ancestral allele indicating that it was neither a constitutively activating nor a null mutation (Fig. 3C). The reintroduction of the Q1197* mutation into MRR1L1191H, yielding MRR1L1191H+Q1197*, resulted in a 128-fold decrease in FLZ MIC (Fig. 3C) and 38-fold lower MDR1 expression values (Fig. 3D) compared to a strain expressing MRR1L1191H and led to a phenotype that mirrored that of MRR1ancestral. Benomyl inducibility of these variants is discussed below.
MRR1Y1126N+P1174P(t) (from U07) and MRR1Y1126N+S359* (from the closely-related U06, Fig. 1A), were similarly analyzed (Fig. 3B). Expression of MRR1Y1126N in U04 mrr1Δ created a strain with a high FLZ MIC (32-64 μg/ml, Fig. 3C) and MDR1 expression (Fig. 3D), similar to that for strains with MRR1Y813C or MRR1L1191H. Addition of the frameshift-inducing indel at P1174, which causes a premature stop codon at N1176 removing 89 amino acids, yielding MRR1Y1126N+P1174P(t), caused a 128-fold decrease in the FLZ MIC and >100-fold decrease in MDR1 expression relative to the strain expressing MRR1Y1126N again leading to a strain that phenocopied that with MRR1ancestral (Fig. 3C and 3D). The addition of the indel at P1174 into an allele with a different constitutively active variant, Mrr1-Y813C, (MRR1Y813C+P1174P(t)) also caused a 256- and >100-fold decrease in FLZ MIC and MDR1 expression, respectively, (Fig. 3C and 3D). In contrast, addition of a SNP causing an early stop codon at S359 to the allele with the activating Y1126N mutation (MRR1Y1126N+S359*) yielded a strain that phenocopied U04 mrr1Δ, indicating this variant was inactive (Fig. 3C and 3D). Together, these data suggest that the Y1126N mutation caused constitutive Mrr1 activity, that was subsequently suppressed by premature stop codons that either restored Mrr1 repression of MDR1 (P1174P(t)) or eliminated activity (S359*). The RNA-Seq analysis supported the results that premature stop codons near the very end of the protein converted constitutively active variants into ones that yielded expression profiles to those for MRR1ancestral and that were distinct from mrr1Δ (Fig. 1D).
In addition to the differences in basal activity, the individual mutations alone and in combination affected chemical inducibility by benomyl. Levels of MDR1 were strongly induced by benomyl in U04 mrr1Δ + MRR1ancestral (40-fold increase), but not in the U04 parental strain with high Mrr1 activity or its mrr1Δ derivative (Fig. 3D). Along with the native Mrr1-Y813C, two other constitutively active Mrr1 variants (Mrr1-L1191H and Mrr1-Y1126N) showed only a 2-3-fold increase in MDR1 expression with benomyl (Fig. 3D) similar to what was observed for more FLZ resistant clinical isolates (Fig. 2D). Surprisingly, addition of the mutations that caused premature stop codons within the last 100 amino acids of Mrr1 to the constitutively active Mrr1-L1191H, Mrr1-Y1126N and Mrr1-Y813C variants restored inducibility by benomyl (Fig. 3D). In fact, there was a strong and significant inverse correlation between basal MDR1 expression and fold induction by benomyl (Fig. 3E).
As in C. albicans, C. lusitaniae Mrr1 regulates the expression of the methylglyoxal reductase encoded by MGD1 (CLUG_01281 or GRP2) (10, 29, 33, 38) and the multidrug efflux pump encoded by FLU1 (CLUG_05825) (10, 39, 40) (Table S1 and Fig. S3A). As with MDR1, expression of both MGD1 and FLU1 was significantly higher in strains encoding the constitutively active Mrr1-Y813C, Mrr1-Y1126N and Mrr1-L1191H variants, compared to a strain encoding the Mrr1-ancestral variant, and the absence of the C-terminus in strains with activating mutations caused a significant decrease in basal MGD1 and FLU1 expression (Fig. S3B and S3C). Benomyl induction of MGD1, like MDR1 (Fig. S3D), was restored upon loss of the C-terminus of the constitutively active Mrr1 variants further supporting the strong negative correlation between basal expression and induction by benomyl (Fig. 2F and S3E). FLU1 expression, however, was not induced by benomyl in any strain suggesting that FLU1 regulation by Mrr1 differs from MGD1 and MDR1 (Fig. 2G and S3F). Interestingly, MDR1 and MGD1, while highly differentially expressed depending on Mrr1 activity (~20-fold or greater), were both de-repressed in the absence of Mrr1, and FLU1 was not and was only weakly differentially expressed (<2-fold) (Fig. 1D and S2B). Together these data indicate the C-terminus of Mrr1 is required for constitutive expression of multiple Mrr1-regulated genes, but not for benomyl induction of the Mrr1-regulated genes tested (Fig. S3A). Combined with the Mrr1 activity across clinical isolates (Fig. 2D), these data indicate that in strains with constitutively active Mrr1 variants, there was selection for mutations to decrease Mrr1 activity, resulting in a mixed population containing constitutively active, truncated but inducible, and loss-of-function Mrr1 variants.
Constitutive Mrr1 activity negatively impacts H2O2 resistance
We next sought to understand why mutations that reduce Mrr1 activity might repeatedly arise in this chronic infection. Previous studies have shown that overexpression of drug efflux pumps in drug resistant microbes can cause a fitness defect due to the energetic cost of constitutive pump production and activity in the absence of a selective substrate (41–43). Deletion of MDR1 from U04 mrr1Δ +MRR1Y813C, which constitutively expresses MDR1, however, did not alter the growth rate across multiple carbon sources (Fig. 4A). In the absence of an obvious fitness defect, we considered factors present in the CF lung, which has been characterized as a highly inflamed environment containing an abundance of neutrophils and macrophages, and high oxidative stress (reviewed in (44, 45)). While little is known about the effects of fungus dominated chronic lung infections in CF, such as the infection from which these isolates originated, an analysis of cytokines within the bronchoalveolar lavage (BAL) fluid from the patient these isolates originated from showed pro-inflammatory cytokines (IL-8 and IL-1β) present were consistent with the neutrophilic environment seen in other patients with CF (Fig. 4B) (45).
In light of these findings, we investigated the effects of Mrr1 activity on reactive oxygen species (ROS) stress generated by hydrogen peroxide (H2O2), a stress strongly associated with high neutrophil counts. In a serial dilution assay, we found that isogenic strains encoding constitutively active Mrr1 variants, while highly resistant to FLZ and diamide (Fig. S4A), had increased sensitivity to 4 mM H2O2 compared to those expressing the Mrr1-ancestral variant (Fig. 4C). Diamide was used to illustrate relative Mrr1 activity instead of FLZ because serial dilution assays on rich medium (YPD) containing FLZ are not always representative of FLZ MIC, which are assessed in defined medium (Fig. S4B). Resistance to H2O2 was restored by addition of mutations causing both mild and severe premature stop codons (Fig. 4C, S4A). The effects of Mrr1 activity on H2O2 sensitivity were independent of strain background, as similar results were seen in isogenic strains in the U04 and U05 backgrounds (Fig. 4C and S4B). Surprisingly, deletion of MDR1 from a strain encoding the constitutively active Mrr1-Y813C variant partially rescued growth (Fig. 4C and S4A), however, the absence of MDR1 did not completely explain the differences as strains lacking MRR1 had increased H2O2 resistance despite elevated MDR1 expression (Fig. 4C). Additionally, the double mutant U04 mrr1Δ mdr1Δ did not have increased resistance to H2O2 compared to U04 mrr1Δ (Fig. 4C), suggesting this may be a complex response. A secondary assay quantifying growth after ~24 hours in liquid cultures containing 1 mM H2O2, though variable day-to-day, confirmed there was a reproducible difference in growth between strains encoding the low activity Mrr1-ancestral and constitutively active Mrr1-Y813C variants (Fig. 4D). Consistent with the plate-based assay, the absence of MDR1 appeared to account for some but not all of the differences in growth in H2O2 (Fig. 4D). To determine if this phenomenon was unique to C. lusitaniae Mrr1 we examined a set of isogenic C. albicans isolates (40), and in vivo or in vitro evolved C. dubliniensis isolates (30) expressing MRR1 alleles containing GOF mutations. We found that for all C. albicans and C. dubliniensis strain sets tested, strains with high Mrr1 activity, which were more resistant to FLZ (40, 46, 47) and diamide, were more sensitive to H2O2 than strains with low Mrr1 activity or lacking MRR1 (Fig. 4C). These data show that the Mrr1 activity driven tradeoff between FLZ and H2O2 resistance is conserved across multiple Candida species.
A screen of isogenic strains for growth in varying concentrations of 48 chemical compounds resuspended from the Biolog Phenotype MicroArrays MicroPlates (Fig. S5) supported our findings that constitutive Mrr1 activity can increase sensitivity to oxidative stress. When comparing strains encoding either the low activity Mrr1-ancestral variant or the constitutively active Mrr1-Y813C variant, with either MDR1 intact or removed, we found there were no differences in growth in the medium used to resuspend the Biolog compounds (Fig. S5A) and many conditions caused less than a 25% difference in growth (Fig. S5B). Unsurprisingly, constitutive Mrr1 activity conferred Mdr1-dependent resistance to twelve compounds, including three triazoles (FLZ, propiconazole, myclobutanil) (Fig. S5B and S5C). High Mrr1 activity also led to Mdr1-independent resistance to four additional compounds, including two other azoles (3-amino-1, 2, 4-triazole and miconazole nitrate) (Fig. S5B and S5C). Eight compounds caused a largely Mdr1-independent decrease in growth in strains encoding the constitutively active Mrr1-Y813C variant: 6-azauracil, berberine, BAPTA, lithium chloride, aminacrine, sodium metasilicate, pentamidine isethionate and potassium chromate (Fig. S5D). Interestingly, berberine and azaserine have previously been studied for their toxic effects on FLZ-resistant Candida strains (48, 49) and calcium inhibitors, such as BAPTA, have been reported to interfere with antifungal resistance (50, 51). While diverse, these compounds are broadly reported to effect metabolism and respiration (52–56), which can lead to oxidative damage via the production of ROS, and/or DNA/RNA integrity, either by direct binding or oxidative damage (57–62). Strain lacking MRR1 or encoding a functional Mrr1 variant that contains a premature stop codon (<100 amino acids removed) were not sensitive to most of these compounds, suggesting secondary mutations causing a decrease or loss of Mrr1 activity could restore resistance in some environments (Fig. S5D).
To gain insight into the mechanisms that lead to differences in oxidative stress resistance between strains with different levels of Mrr1 activity, we compared the gene expression profiles after a 30-minute exposure to 0.5 mM H2O2, a partially inhibitory concentration. H2O2 exposure had broad strain-independent effects on the transcriptome, altering expression of 786 genes (FC≥2, FR<0.05) including increased expression of CLUG_04072, a homolog of C. albicans CAT1, which was previously shown to be important for the resistance of C. lusitaniae to H2O2 (63) (Fig. S6A and Table S2). While there were subtle differences in the H2O2 response between strains expressing the constitutively active Mrr1-Y813C variant compared to U04 mrr1Δ MRR1ancestral there were no clear patterns that would explain the difference in H2O2 resistance (Fig. S6B). The majority of differences in gene expression were seen in the magnitude of induction of Mrr1-regulated genes by H2O2, a known inducer of Mrr1 in other species (29, 64), indicating that, as with benomyl (Fig. S3D-F), strains with constitutively active Mrr1variants are less inducible than strains with low activity variants (Fig. S6B and S6C). Next, we investigated the expression of homologs of oxidative stress response (OSR) genes previously characterized in C. albicans and S. cerevisiae and found that there was not a significant Mrr1-dependent difference in basal or H2O2-induced expression of these genes (Fig. S6A). Genes assessed included the oxidative stress responsive transcription factor encoded by CaCAP1 or ScYAP1, superoxide dismutase (SOD2, SOD4, SOD6), enzymes involved in the thioredoxin (TSA1, TRX1, TRR1) and glutathione (GPX, GSH1) systems, catalase (CAT1), and OSR genes involved in carbohydrate metabolism and the DNA-damage response (65, 66). Further analysis is required to better understand the link between constitutive Mrr1 activity and H2O2 sensitivity, however these data highlight that the sensitivity is not due to failure to induce an oxidative stress response, but more-likely a consequence of the activity of Mrr1-regulated genes, such as MDR1 (Fig. S4A and S4C).
Phenotype dynamics in chronic infection populations over time
In light of the evidence for complex evolution of MRR1 and the potentially advantageous phenotypes associated with both high and low Mrr1 activity, we sought to better understand the fractions of isolates with these Mrr1 associated traits over time. For this analysis, we used arrayed C. lusitaniae populations isolated from sputum or one BAL procedure collected from the same subject over three years, with the first time point approximately six months after the first clinical culture report of the high levels of “non-albicans Candida” (NAC) as shown in Fig. 5A. Upon plating isolates on agar with FLZ (8 μg/ml) or H2O2 (4 mM) (Fig. 5A), we found an inverse correlation between robust growth on FLZ and robust growth on H2O2. It was uncommon for isolates to be inhibited or uninhibited in both conditions (Fig. S5A). Isolates from the early samples were predominately sensitive to FLZ (10), but were largely resistant to H2O2. During and soon after the course of FLZ therapy (Sp1.5 and Sp2, respectively), however, there was an increase isolates that were more FLZ resistant but H2O2-sensitive (Fig. 5A). Subsequent samples from two years after the FLZ therapy was completed varied in the proportion of isolates that grew better on H2O2 and FLZ. Thus, the C. lusitaniae population shifted back and forth between being dominated by isolates with higher H2O2 resistance or higher FLZ resistance, but both phenotypes remained in the population over time (Fig. 5B).
Discussion
A population of C. lusitaniae isolates first described in Demers et al. (10) contained an unexpectedly large number of nonsynonymous mutations in the gene encoding the transcription factor, Mrr1, which regulated FLZ resistance, suggesting that Mrr1 activity was under strong selective pressure in vivo. These MRR1 alleles contained either one or two nonsynonymous SNPs or indels (Fig. 1A) and isolates with one mutation had on average higher FLZ resistance than those with two nonsynonymous MRR1 mutations (Fig. 2A). While multiple studies have shown that constitutive Mrr1 activity is beneficial under multiple biologically relevant conditions, including exposure to azoles (10, 29), bacterial-produced toxins including phenazines (10), and host-produced antifungal peptides including histatin 5 (10, 40), it was unclear why MRR1 alleles conferring low Mrr1 activity would be selected for in this population (10). Deconstruction of MRR1 alleles with two mutations revealed an evolutionary path on which an activating mutation arose first, followed by suppressing mutations that either restored low basal activity but retained inducibility, or abolished Mrr1 activity altogether (Fig. 3 and S3). Interestingly, a C. parapsilosis strain was recently found to contain a central domain mutation and a C-terminal truncation (Mrr1P295L+Q1074*) similar to the alleles described above, however, it is not currently known how these mutation impact Mrr1 activity and FLZ resistance (67), suggesting that selection for and against elevated Mrr1 activity may also occur in other Candida species.
Surprisingly, the RNA-Seq analysis of isogenic strains expressing different MRR1 alleles revealed that C. lusitaniae Mrr1 appears to positively and negatively regulate genes expression (Fig. 1D) although further analysis is required to determine which genes are direct targets of Mrr1. Adding to previous studies in C. lusitaniae (10, 13) and C. albicans (29), we found that Mrr1 positively regulates 41 genes with a fold change ≥ 2 and 102 genes with a fold change ≥ 1.5 (FDR<0.05). Mrr1-induced genes include multiple MFS and ABC transporters (i.e. MDR1, FLU1, CDR1), methylglyoxal reductases (33), putative alcohol dehydrogenases, and a variety of other putative metabolic genes (Table S1). Constitutively active Mrr1 also appears to repress expression of 42 genes (fold change ≥ 1.5, FDR<0.05), including multiple iron and/or copper transporters and reductases, and sugar transporters (Table S1). These data combined with Bierman et al., which showed that C. lusitaniae Mrr1 is induced by the spontaneously formed stress signal methylglyoxal (33), imply that Mrr1 may play a larger role in a generalized metabolic or stress response, beyond what has been previously studied in response to FLZ and xenobiotic stressors.
While the C-terminal region of C. lusitaniae Mrr1 was necessary for constitutive Mrr1 activity, it was not required for induction of Mrr1-regulated genes, including MDR1 and MGD1, in response to benomyl (Fig. 2D, 3 and S3). Subsequent addition of mutations resulting in the loss >200 amino acids, however, caused a slight decrease in FLZ resistance and MDR1 expression, but these variants were no longer inducible by benomyl and phenocopied strains completely lacking MRR1 (Fig. S1B, 3C and 3D). These data are consistent with previous studies showing C-terminal truncations prior to amino acid 944 in C. albicans MRR1, homologous to position 1116 in C. lusitaniae MRR1, caused a complete loss of CaMrr1 activity (68). The L11, L12 and U06 strains encoding Mrr1 variants with premature stop codons before amino acid 1116 similarly phenocopied mrr1Δ strains (Fig. 2C and 2D). Surprisingly, loss-of-function Mrr1 variants and mrr1Δ strains had intermediate expression of a subset of the most strongly differentially regulated genes compared to strains with low activity Mrr1 (Fig. S2B), which has not been observed in other Candida species (29, 31, 32). Additional studies are required to determine if this phenomenon is unique to C. lusitaniae or more broadly shared among non-albicans Candida species closely related to C. lusitaniae, such as C. auris (20, 26), and if any of the co-regulators of the Mrr1 regulon described in C. albicans (64, 69, 70) are involved. These findings raised the question as to why, if constitutive Mrr1 was initially selected for, would it later be selected against in vivo, especially in the absence of an obvious growth defect (Fig. 4A and S5A).
Chronic lung infections are typically an inflamed environment (Fig. 4B) containing a high number of polymorphonuclear leukocytes (PMNs) that produce proteases, myeloperoxidases and ROS (71, 72), which is an important component of the immune system used to kill fungi (reviewed in (73)). In a screen of diverse chemical compounds, we found that strains with constitutive Mrr1 activity were more strongly inhibited by multiple compounds that have previously been shown to cause damage through oxidative stress (Fig. S5B-D). When we specifically interrogated H2O2 resistance, we found that C. lusitaniae strains encoding constitutively active Mrr1 variants were more sensitive than strain encoding low activity Mrr1 variants or lacking a functional Mrr1 (Fig. 4A, S4). Sensitivity to H2O2 and the compounds from the Biolog plates was at least partially dependent on Mdr1, thought other Mrr1-regulated genes may still contribute to the decreased growth under conditions of oxidative stress (Fig. S4B, S5 and S6). Interestingly, the tradeoff between FLZ and H2O2 resistance was conserved broadly among a time series of C. lusitaniae isolates and other Candida species (Fig. 4C and 5A).
As outlined in the model in Figure 5B, together these data highlight that changing environments within complex and dynamic chronic infections could contribute to the development of heterogeneous fungal populations. Though it appears that initial selection on the ancestral version of Mrr1 was driven by the need for increased Mrr1 activity, over time either these selective pressures were removed, or other pressures became dominant, resulting in a secondary wave of mutations. This secondary wave of mutations caused a decrease or loss of Mrr1 activity, which uniquely to C. lusitaniae further contributed towards a population with mixed levels of FLZ resistance (Fig. 5B). Though the exact selective pressures at play in this instance are unknown, these data highlight the importance of understanding how microbes evolve in vivo, as complex environments, even in the absence of clinically used antifungals, can shape the microbial population and lead to antimicrobial resistance.
Materials and Methods
Strains and growth conditions
Candida strains used in this study are listed in Table S3. All strains were stored as frozen stocks with 25% glycerol at −80 °C and subcultured on YPD (1% yeast extract, 2% peptone, 2% glucose, 1.5% agar) plates at 30 °C. Strains were regularly grown in YPD liquid medium at 30 °C on a roller drum. Cells were grown in YNB (0.67% yeast nitrogen base medium with ammonium sulfate (RPI Corp)) liquid supplemented with either 2% glucose, 2% glycerol or 2% casamino acids and in RPMI-1640 (Sigma, containing L-glutamine, 165 mM MOPS, 2% glucose) liquid as noted. Media was supplemented with 8 μg/ml FLZ (stock 4 mg/ml in DMSO), 1 mM diamide (stock 58 mM in water) or 1-6 mM H2O2 (30% w/v in water, ~9.8M) as noted. Escherichia coli strains were grown in LB with either 100 μg/ml carbenicillin or 15 μg/ml gentamycin as necessary to obtain plasmids. BAL fluid and sputum were obtained in accordance with institutional review board protocols as described in (74).
DNA for gene knockout constructs
Gene replacement constructs for knocking out MRR1 (CLUG_00542, as annotated in (10)) and MDR1 (CLUG_01938/9 (10)) were generated by fusion PCR as described in Grahl et al. (63). All primers (IDT) used are listed in Table S4. Briefly, 0.5 to 1.0 kb of the 5’ and 3’ regions flanking the gene was amplified from U04 DNA, isolated using the MasterPure Yeast DNA Purification Kit (epiCentre). The nourseothricin (NAT1) or hygromycin (HygB) resistance cassette was amplified from plasmids pNAT (75) and pYM70 (76), respectively, using the Zyppy Plasmid Miniprep kit (Zymo Research). Nested primers within the amplified flanking regions were used to stitch the flanks and resistance cassette together. PCR products for transformation were purified and concentrated with the Zymo DNA Clean & Concentrator kit (Zymo Research) with a final elution in molecular biology grade water (Corning).
DNA for insertion of NAT1 at neutral site in C. lusitaniae genome
The approximately 4000 bp genomic region between CLUG_03302 and CLUG_03303 on chromosome 4, which was not predicted to contain any genes or promoter regions, was targeted as a potentially neutral insertions site. To create plasmid DH3261 containing NAT1 flanked by homology to this region of chromosome 4, approximately 1.0 kb of the flanking regions (positions 228,652 – 229,651 and 229,701 – 230,691) were amplified from U05 gDNA. All primers (IDT) used are listed in Table S4. NAT1was amplified from pNAT (75). PCR products were purified and concentrated then assembled with the vector (pRS426 (77) linearized with KpnI-HF and SalI-HF (New England BioLabs) and treated with the phosphatase rSAP (New England BioLabs)) using the NEBuilder HiFi DNA Assembly cloning kit (New England BioLabs). Assemblies were transformed into High Efficiency NEB®5-alpha competent E. coli (New England BioLabs). The NAT1 insertion construct was isolated from DH3261 by digestion with KpnI-HF and SalI-HF (New England BioLabs).
Plasmids for complementation of MRR1
Plasmids for complementing MRR1 were created as described in Biermann et al, 2020 (33). For naturally occurring MRR1 alleles, we amplified i) the MRR1 gene and terminator with ~1150 bp upstream for homology from the appropriate strain’s genomic DNA, ii) the selective marker, HygB from pYM70 (76), and iii) ~950 bp downstream of MRR1 for homology from genomic U05 (identical sequence for all relevant strains) using primers (IDT) listed in Table S4. PCR products were cleaned up using the Zymo DNA Clean & Concentrator kit (Zymo Research) and assembled using the S. cerevisiae recombination technique previously described (78). Plasmids created in S. cerevisiae were isolated using a yeast plasmid miniprep kit (Zymo Research) and transformed into High Efficiency NEB®5-alpha competent E. coli (New England BioLabs). E. coli containing pMQ30 derived plasmids were selected for on LB containing 15 μg/ml gentamycin. Plasmids from E. coli were isolated using a Zyppy Plasmid Miniprep kit (Zymo Research) and subsequently verified by Sanger sequencing with the Dartmouth College Genomics and Molecular Biology Shared Resources Core. MRR1 complementation plasmids were linearized with Not1-HF (New England BioLabs), cleaned up the Zymo DNA Clean & Concentrator kit (Zymo Research) and eluted in molecular biology grade water (Corning) before transformation of 2 μg into C. lusitaniae strain U04 mrr1Δ as described below.
The MRR1ancestral allele sequence was amplified from gDNA of a closely related C. lusitaniae isolate that had the same MRR1 sequence but lacked any of the nonsynonymous mutation that varied among the population of C. lusitaniae isolates described here. This MRR1 sequences does contain multiple synonymous and nonsynonymous mutations in comparison with that of the reference strains, ATCC 42720 (79). Additional MRR1 alleles were amplified from gDNA from U04 (MRR1Y813C), U05 (MRR1L1191H+Q1197*), U02 (MRRY1126N+P1174P(t)) and U06 (MRR1S359*+Y1126N). While making the pMQ30MRR1-S359*+Y1126N plasmid, one clone was identified that lacked the S359* mutation resulting in the pMQ30MRR1-Y1126N plasmid. To create additional MRR1 alleles that were not identified within any C. lusitaniae isolates, pieces of MRR1 were selectively removed and repaired with DNA either containing or lacking the desired mutation. Because the L1191H and Q1197* mutations were so close together, an alternate strategy was used to separate these mutations from the MRR1L1191H+Q1197* allele. DNA fragments synthesized by IDT containing either the L1191H or Q1197* mutations alone (sequences in Table S4) were amplified then assembled with pMQ30MRR1-L1191H+Q1197* (linearized with PvuI-HF) using the NEBuilder HiFi DNA Assembly cloning kit (New England BioLabs). To remove an unexpected nonsynonymous mutation in pMQ30MRR1-Q1197*, this plasmid was digesting with EcoNI and repaired with a piece of DNA amplified from U04 mrr1Δ+MRR1ancestral lacking the unwanted mutation. pMQ30MRR1 complementation plasmids was digested with Not1-HF (New England BioLabs) for transformation.
Strain construction
Mutants were constructed as previously described in Grahl et al. using an expression free ribonucleoprotein CRISPR-Cas9 method (63). 1 to 2 μg of DNA for gene knockout constructs generated by PCR or 2 μg of digested plasmid, purified and concentrated with a final elution in molecular biology grade water (Corning), was used per transformation. Plasmids containing complementation and knockout constructs and resulting strains are listed in Table S3 and crRNAs (IDT) are listed in Table S4. Transformants were selected on YPD agar containing 200 μg/mL nourseothricin or 600 μg/mL hygromycin B.
Drug susceptibility assays
Minimum inhibitory concentration (MIC) was determined using a broth microdilution method as previously described (80) with slight modifications (10). Briefly, 2×103 cells were added to a two-fold dilution series of FLZ prepared in RPMI-1640, starting at an initial concentration of 64 μg/ml, then incubated at 35 °C for 24 hours. The MIC was defined as the drug concentration that abolished visible growth compared to a drug-free control.
Quantitative RT-PCR
Overnight cultures were back diluted to an OD600 of ~0.1 and grown for 6 hours in YPD liquid medium at 30°C. 50 μg/ml of benomyl (stock 10 mg/ml in DMSO) or an equivalent volume of DMSO were added for experiments assessing the induction of Mrr1 activity. 7.5 μg RNA (harvested using the MasterPure Yeast RNA Purification Kit (Epicentre)) was DNAse treated with the Turbo DNA-free Kit (Invitrogen). cDNA was synthesized from 300-500 ng of DNAse-treated RNA using the RevertAid H Minus First Strand cDNA Synthesis Kit (Thermo Scientific), following the manufacturer’s instructions for random hexamer primer (IDT) and GC rich template. qRT-PCR was performed on a CFX96 Real-Time System (Bio-Rad), using SsoFast Evergreen Supermix (Bio-Rad) with the primers listed in Table S4. Thermocycler conditions were as follows: 95 °C for 30 s, 40 cycles of 95 °C for 5 s, 65 °C for 3 s and 95 °C for 5 s. Transcripts were normalized to ACT1 expression.
RNA Sequencing
Overnight cultures were back diluted into YPD and grown to exponential (~8 h) twice, then treated with vehicle or 0.5 mM H2O2 for 30 minutes, in biological triplicate. RNA was harvested from snap-frozen pellets (using liquid nitrogen) using the MasterPure Yeast RNA Purification Kit (Epicentre) and stored at −80 °C. RNA libraries were prepared using the Kapa mRNA HyperPrep kit (Roche) and sequenced using single-end 75 bp reads on the Illumina NextSeq500 platform. The data analysis pipeline is available in github repository (https://github.com/stajichlab/RNASeq_Clusitaniae_MRR1) and archived as DOI: [To be generated]. FASTQ files were aligned to the ATCC 42720 (79) genome with the splice-site aware and SNP tolerant short read aligner GSNAP (v v2019-09-12) (81). The alignments were converted to sorted BAM files with Picard (v2.18.3; https://broadinstitute.github.io/picard/) and read counts computed with featureCounts (v1.6.2) (82) with updated genome annotation to correct truncated gene model for locus CLUG_00542, and combine a single gene split into two, CLUG_01938_1939; reasoning for these changes explained in (10). Differential gene expression analyses were performed with the edgeR (83) package in Bioconductor, by fitting a negative binomial linear model. The resulting P values were corrected for multiple testing with Benjamini-Hochberg to control the false discovery rate. Genes for which there were less than 2 counts per million (CPM) across the three (absent genes) were not included for differentially expressed gene analysis. Two separate linear models were created to define the Mrr1 regulon in control conditions alone and determine the interaction between Mrr1 activity and H2O2 exposure.
To define the Mrr1 regulon in YPD alone we identified genes differentially expressed between strains with constitutive Mrr1 activity (U04 and U04 mrr1Δ +MRR1Y813C) and low Mrr1 activity (U04 mrr1Δ, U04 mrr1Δ +MRR1ancestral, and U04 mrr1Δ +MRR1L1191H+Q1197*); this model contained 5,474 genes. We discarded genes for which i) the log2FC greater than 1 (2-fold, see Fig. 1B) or 0.585 (1.5-fold, see Table S1) with an FDR<0.05, (ii) the average CPMs for replicates was not greater than 10 for any strain, and ii) expression in both U04 and U04 mrr1Δ +MRR1Y813C was similar. Results are summarized in Table S1, including the Mrr1 regulon (Table S1a), and the normalized CPMs/gene used for this linear model(Table S1b).
To determine how constitutive Mrr1 activity impacted the response to H2O2 we identified the overlap between the interaction between U04 or U04 mrr1Δ +MRR1Y813C and exposure to 0.5 mM H2O2, as compared to the reference strain (U04 mrr1Δ +MRR1ancestral) and condition (YPD alone); this model contained 5600 genes. Results are summarized in Table S2, including the interaction between strains with constitutively active Mrr1 and H2O2 (Table S2a), the effect of H2O2 treatment (Table S2b), and all normalized CPMs/gene used for this linear model (Table S2c).
Biolog Phenotype MicroArrays analysis
For the chemical sensitivity screen, the chemicals in Biolog plates PM22D and PM24C were resuspended in 100 ul YPD liquid and transferred to a sterile 96-well polystyrene plate (Fisher) for kinetic measurements. 100 ul of cells adjusted to an OD of 0.01 in YPD was added to each well. Plates were incubated at 37 °C for 24 hours. A control plate containing no drug was grown simultaneously for comparison.
Serial dilution assays
Following growth in YPD medium overnight with aeration at 30°C, cultures were diluted in water to an OD600 of 1. Serial dilutions of ten-fold were carried out in a microtiter plate to yield six concentrations ranging from approximately 107 cells/ml (for OD600 of 1) to approximately 102 cells/ml. 5 μl of each dilution were applied to YPD plates containing 4 or 5 mM H2O2 (stock 30% w/v, 9.8 M) or 1 mM diamide (stock 58 mM in water). Images were captured after incubation at 37°C for 24 or 48 hours.
Luminex Analysis
Cytokines in BAL fluid samples were measured (pg/ml) in singlicate by Luminex using a Millipore human cytokine multiplex kits (EMD Millipore Corporation, Billerica, MA) according to manufactures instructions. Assays were performed by the DartLab – Immune Monitoring and Flow Cytometry Resource core at Dartmouth.
Statistical analyses
Statistical analyses were done using GraphPad Prism 6 (GraphPad Software). Unpaired Student’s t-tests (two-tailed) with Welch’s correction were used to evaluate the difference in FLZ MIC between isolates containing one of two mutations in MRR1. One and two-way ANOVA tests were performed across multiple samples with either Tukey’s multiple comparison test for unpaired analyses or Sidak’s multiple comparison test for paired analyses conducted in a pairwise fashion. P values <0.05 were considered as significant for all analyses performed and are indicated with asterisks: *P<0.05, **P<0.01, ***P<0.001 and ****P<0.0001.
Data availability
The data supporting the findings in this study are available within the paper and its supplemental information and are also available from the corresponding author upon request. The raw sequence reads from the RNA-Seq analysis have been deposited into NCBI sequence read archive under BioProject PRJNA680763. Raw and processed RNA-Seq count data are available in Gene Expression Omnibus (GSE162151) and include minor updates to the genome annotation and assembly for C. lusitaniae.
Acknowledgments
We would like to thank J. Morschhäuser (Universität Würzburg) and L. Myers (Dartmouth College) for sharing C. albicans and C. dubliniensis strains. Research reported in this publication was supported by National Institute of Health (NIH) grant R01 AI127548 to D.A.H. from the National Institute of Allergy and Infectious Disease, R01 HL122372 to A.A. from the National Heart, Lung and Blood Institute, and National Institute of General Medical Sciences (NIGMS) of the NIH under award number T32GM008704 and AI133956 to E.G.D. J.E.S. is a CIFAR Fellow in the program Fungal Kingdom: Threats and Opportunities. This work was also supported by the Cystic Fibrosis Foundation Research Development Program (CFFRDP) STANTO19R0 for the Translational Research Core, and ASHARE20P0 to A.A. Sequencing services and specialized equipment was provided by the Genomics and Molecular Biology Shared Resource Core at Dartmouth, and Luminex analysis was performed by the DartLab – Immune Monitoring and Flow Cytometry Resource Core at Dartmouth, both supported by NCI Cancer Center Support Grant 5P30CA023108-37. Equipment used was supported by the NIH IDeA award to Dartmouth BioMT P20-GM113132. Analyses were performed using the computational and data storage resources of the University of California-Riverside HPCC funded by grants from the National Science Foundation (NSF) (MRI-1429826) and NIH (1S10OD016290-01A1). The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH.
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