Expanding the toolkit for genetic manipulation and discovery in Candida species using a CRISPR ribonucleoprotein-based approach

Abstract

The World Health Organization recently published the first list of priority fungal pathogens highlighting multiple Candida species including C. glabrata, C. albicans, and C. auris. The use of CRISPR-Cas9 and auxotrophic C. glabrata and C. albicans strains have been instrumental in the study of these fungal pathogens. Dominant drug resistance cassettes are also critical for genetic manipulation and eliminate the concern of altered virulence when using auxotrophic strains. However, genetic manipulation has been mainly limited to the use of two drug resistance cassettes, NatMX and HphMX. Using an in vitro assembled CRISPR-Cas9 ribonucleoprotein (RNP)-based system and 130–150 bp homology regions for directed repair, we expand the drug resistance cassettes for Candida to include KanMX and BleMX, commonly used in S. cerevisiae. As a proof of principle, we demonstrated efficient deletion of ERG genes using KanMX and BleMX. We also showed the utility of the CRISPR-Cas9 RNP system for generating double deletions of genes in the ergosterol pathway and endogenous epitope tagging of ERG genes using an existing KanMX cassette. This indicates that CRISPR-Cas9 RNP can be used to repurpose the S. cerevisiae toolkit. Furthermore, we demonstrated that this method is effective at deleting ERG3 in C. auris using a codon optimized BleMX cassette and effective at deleting the epigenetic factor, SET1, in C. albicans using a recyclable SAT1. Using this expanded toolkit, we discovered new insights into fungal biology and drug resistance.

INTRODUCTION

Fungal infections pose a significant public health concern, with over a billion superficial infections and 1.5 million deaths occurring annually worldwide (1, 2). Candida species are responsible for roughly 40–70% of invasive fungal infections (1–3), and several species are classified as “high priority fungal pathogens” by the World Health Organization (WHO) for study, including C. glabrata, C. albicans, and C. auris. Infections can range from superficial to life-threatening, with invasive candidiasis leading to a mortality rate of 20–60% (4, 5). Currently, there are three major antifungals clinically used for treatment of fungal infections; azoles, echinocandins, and polyenes (6–8). However, antifungal drug resistance has become a significant concern, highlighted by the increase in clinically acquired drug resistance in C. albicans and C. glabrata and the recent emergence of a multi-drug resistant pathogen, C. auris (8, 9). With increased drug resistance and emerging pathogens, there is an urgent need for the development and expansion of new and existing tools for studying drug resistance and pathogenesis in Candida, especially in non-albicans Candida (NAC) species.

To address this need, several groups use various Clustered Regular Interspaced Short Palindromic Repeats (CRISPR) based methods for genetic manipulations. CRISPR-Cas9 is a tool that utilizes the Cas9 endonuclease to direct double stranded breaks (DSBs) at the desired locus by binding to a gene specific guide RNA followed by a protospacer-adjacent motif (PAM) sequence. A Cas9-mediated DSB will activate either the nonhomologous end-joining (NHEJ) for error-prone repair resulting in insertions or deletions or a precise homology directed repair (HDR) using a donor template. Using both approaches can greatly enhance the efficiency to generate genetic mutations, gene replacements, or epitope tags.

There are two common strategies for utilizing CRISPR genome editing in C. albicans or C. glabrata. One approach involves the expression of the Cas9 enzyme and sgRNA from separate plasmids while the other approach uses one plasmid for expressing Cas9 and sgRNA (10, 11). These plasmid-based approaches can be either episomally expressed or integrated in the genome (12, 13). Another approach is using an expression-free CRISPR-Cas9 ribonucleoprotein (RNP) method (10, 14–18). The CRISPR-Cas9 RNP approach has been used with a HDR template containing the NAT1, SAT, and HygB resistances cassette for generating gene deletions in C. glabrata, C. auris, C. lusitaniae, and C. albicans (14–18). A major advantage of this system is that the CRISPR-based RNP system does not require plasmid engineering or species-specific promoter expression in cells (10, 11, 18). Instead, recombinant Cas9 protein, crRNA and tracrRNA are assembled as a ribonucleoprotein complex in vitro and electroporated into competent cells which reduces the steps needed to genetically manipulate prototrophic strains or clinical isolates.

In this study, we used an expression-free CRISPR-Cas9 RNP-based approach using homology regions of 130–150 bp for making efficient gene deletions in C. glabrata and C. auris. Our CRISPR-Cas9 RNP approach showed improved efficiency over a non-CRISPR based method using ADE2 as our reporter for gene disruption. Furthermore, all drug resistance cassettes used for gene disruptions and epitope tagging were PCR amplified using homology regions of 130–150 bp, indicating large flanking sequences are not required with the CRISPR-Cas9 RNP-based approach. Our approach also permits making double deletions and epitope tagging which are difficult to make without CRISPR. More importantly, we demonstrated the utilization of drug resistance cassettes KanMX and BleMX in C. glabrata for generating gene deletions and KanMX for generating epitope tags. These two drug resistance cassettes have not been widely used for C. glabrata but are extensively used for S. cerevisiae. Finally, we demonstrated that the CRISPR-Cas9 RNP approach can also be utilized for making gene deletions in C. auris using codon optimized BleMX. Overall, using the CRISPR-Cas9 RNP approach allowed us to expand the fungal pathogen toolkit by demonstrating that KanMX containing plasmids used for S. cerevisiae can be repurposed for C. glabrata and that BleMX can be used for C. auris and C. glabrata. Furthermore, we showed the utility of these tools by providing phenotypic characterization of factors that alter ergosterol biosynthesis and fluconazole drug susceptibility.

RESULTS

The CRISPR-Cas9 RNP system for efficient gene replacement in C. glabrata using 130–150 bp homology regions.

CRISPR-mediated or non-CRISPR-based methods generally rely on large flanking homologous regions ranging from 500 bp to 1000 bp for efficient gene replacements in Candida species (18, 19). Often steps to generate long flanking regions are time consuming and tedious using either cloning or multi-step fusion PCR approaches. The initial CRISPR-Cas9 RNP system developed for Candida species including C. glabrata utilized long homology regions ranging from 500 to 1000 bp (18). However, using CRISPR-Cas9 plasmid-based system and auxotrophic cassettes, it has been reported for C. glabrata that flanking homology regions ranging from 20–200 bp can be used for gene insertions resulting in gene disruption (12). To determine if short homology regions flanking drug resistance cassettes were efficient in making gene deletions in C. glabrata using a CRISPR-Cas9 RNP method, we PCR amplified drug resistance cassettes using long oligonucleotides (IDT Ultramers) that range from ~130–150 bp of homology to the ADE2 gene. The ADE2 gene was selected due to its red pigment phenotype when ADE2 gene is disrupted which allows for quick determination of gene replacement efficiency and has been commonly used to determine CRISPR efficiency (13, 20, 21). Using the pAG25 NatMX and pAG32 HphMX plasmids (Fig. 1A, 1B) (22), we deleted the entire ADE2 open reading frame and counted the proportion of white and red colonies (Fig. 1C). With the addition of a CRISPR-Cas9 RNP containing two gRNAs and 130–150 bp of flanking homology, we observed a five-fold increase in the proportion of red colonies compared to the cassette alone (Fig. 1D). With an efficiency of 62% red colonies, we determined that long homology regions are not required for efficient gene replacement in C. glabrata using NatMX. Similarly, we observed a five-fold increase in the proportion of red colonies using Hygromycin B (HphMX), with an efficiency of 55% replacement using our CRISPR-Cas9 RNP method (Fig. 1E). Altogether, these data suggest that our modified CRISPR-RNP method using 130–150 bps of homology can efficiently generate single gene deletions in C. glabrata.

FIG 1.

The CRISPR-Cas9 RNP system using 130–150 bp homology regions efficiently generates ADE2 deletions in C. glabrata using NatMX and HphMX. (A) Schematic of pAG25 NatMX plasmid. P1 and P2 indicate location of amplification sequences. (B) Schematic of pAG32 HphMX plasmid. P1 and P2 indicate location of amplification sequences. (C) Representative transformation plate for ADE2 deletion using NatMX with and without addition of CRISPR-RNP. (D) Total number of positive transformants using NatMX with and without addition of CRISPR-RNP. Numbers represent the summation across three separate transformations. (E) Total number of positive transformants using HphMX with and without addition of CRISPR-RNP. Numbers represent the summation across three separate transformations.

Using CRISPR-Cas9 RNP system to generate sequential gene replacements utilizing NatMX and HphMX resistance cassettes.

After determining this system efficiently generates single gene deletions, we then tested whether the CRISPR-Cas9 RNP method was sufficient for making sequential gene disruptions for generating double deletion strains. While the pAG25 and pAG32 plasmids are effective for use in single deletions, it is often difficult to generate double gene deletions with these replacement cassettes using standard transformation methods. We suspect that making double deletions using drug resistance cassettes are difficult because they often share homology with the TEF1 promoter and TEF1 terminator (Fig. 1A, 1B). To overcome these issues, we tested if our CRISPR-Cas9 RNP method was sufficient for generating double gene deletions using HphMX and NatMX resistance cassettes.

For proof of principle, we probed the ergosterol biosynthesis pathway, a critical pathway for azole antifungal drugs. Azole drugs inhibit Erg11, lanosterol 14-alpha-demethylase, to block ergosterol biosynthesis and leads to accumulation of an Erg3-dependent toxic sterol 14α-methyl-3,6-diol and growth arrest (23–25). Thus, ERG gene deletions or mutations in this pathway can alter azole susceptibility and growth. For example, ERG3 is known to have an azole resistant phenotype when deleted or mutated in S. cerevisiae or C. albicans which is a consequence of not producing the toxic sterol 14α-methyl-3,6-diol (24, 26, 27). However, there has been conflicting results in C. glabrata where ERG3 deletions do not confer resistance to fluconazole while microevolved ERG3 mutations and clinical isolates have shown resistance to fluconazole (25, 28–31). To address this issue, we used our CRISPR-Cas9 RNP method to make erg3Δ strains using the pAG25-NatMX and pAG32-HphMX as a template or erg5Δ strain using NatMX (Fig. 1A, B). We performed five-fold serial dilution spot assays on these strains to confirm and compare their phenotypes with and without 64 μg/mL fluconazole in SC media (Fig. 2A, B). Both erg3Δ strains demonstrate a slow growth phenotype, but also a clear increased resistance to fluconazole (Fig. 2A). However, the erg5Δ strain did not have an observable growth defect without fluconazole and little to no growth on fluconazole containing plates (Fig. 2B).

FIG 2.

The CRISPR-Cas9 RNP system generates single and double gene deletions utilizing NatMX and HphMX in C. glabrata. (A and B) Five-fold serial dilution spot assays with and without 64 μg/mL fluconazole (FLZ). Indicated single deletion strains were generated using the CRISPR-Cas9 RNP method. Double deletion strains were generated using CRISPR-Cas9 RNP method sequentially and three independent clones are shown. Images were captured at 48 hours. (C and D) Expression of the indicated genes were determined by qRT-PCR analysis of mid-log phase cells. Data was normalized to RDN18 mRNA levels and are the average of three biological replicates with three technical replicates each. Error bars represent the standard deviation.

Next, we used our CRISPR-Cas9 RNP method to generate erg3Δerg5Δ double deletion strains, by deleting ERG5 with HphMX in the previously constructed erg3Δ (NatMX) strain. After confirming positive transformants via colony PCR, five-fold serial dilution spot assays with and without 64 μg/mL fluconazole were performed. Interestingly, all erg3Δerg5Δ strains display a synthetic growth defect, more than what was observed in the single erg3Δ and erg5Δ strains (Fig. 2B). Despite this significant growth defect under untreated conditions, erg3Δerg5Δ strains were still able to grow on fluconazole containing plates similar to an erg3Δ strain (Fig. 2B). To further validate these strains, we grew cells in SC media to mid-log phase and collected cells for qRT-PCR expression on both ERG3 and ERG5. In each strain lacking ERG3, we detected no ERG3 transcript, confirming that ERG3 was deleted (Fig. 2C). Additionally, in each strain lacking ERG5, we detected no ERG5 expression (Fig. 2D), confirming ERG5 was deleted. Interestingly, we do see decreased expression of ERG3 in the erg5Δ strain and increased expression of ERG5 in the erg3Δ strain which is consistent with what is observed in S. cerevisiae (32, 33). Altogether, these results suggest that our CRISPR-Cas9 RNP method permits engineering of single and double deletions in C. glabrata. Moreover, we clearly established that erg3Δ strains are resistant to fluconazole and identify a genetic interaction between ERG3 and ERG5.

The CRISPR-Cas9 RNP system efficiently generates gene deletions utilizing the BleMX drug resistance cassette in C. glabrata.

Since our CRISPR-Cas9 RNP system is efficient at generating single and double deletions in C. glabrata using NatMX and HphMX, we then tested whether this system was effective for using other drug resistance cassettes typically not used in C. glabrata. We first tested BleMX, which confers resistance to Zeocin, as the use of BleMX has been reported once in C. glabrata using a non-CRISPR transformation method, albeit at extremely low efficiency (<1%) (34). To first determine whether the CRISPR-Cas9 RNP system effectively generates gene deletions using BleMX, we deleted the entire open reading frame of ADE2 using the pCY3090-07 plasmid as a template (Fig. 3A) (35). When comparing the proportion of red colonies with and without the addition of CRISPR-Cas9, a five to six-fold increase in efficiency was observed (Fig. 3B). Next, using the pCY3090-07 plasmid, we deleted ERG3 using our CRISPR-Cas9 RNP method and subsequently performed a five-fold serial dilution spot assay with and without 64 μg/mL fluconazole to compare phenotypes of these strains with previously constructed erg3Δ strains. Similar to the previously constructed erg3Δ strains, we again observed an azole resistant phenotype. Thus, our results using 4 different drug resistance cassettes clearly indicate that erg3Δ strains are resistant to fluconazole under the indicated conditions (Fig. 3C and 4C). These data show that BleMX is an effective dominant drug resistance cassette in C. glabrata when using the CRISPR-Cas9 RNP system.

FIG 3.

The CRISPR-Cas9 RNP system efficiently generates gene deletions utilizing BleMX in C. glabrata. (A) Schematic of pCY3090-07 plasmid. P1 and P2 indicate location of amplification primer sequences. (B) Total number of positive transformants using BleMX with and without addition of CRISPR-Cas9 RNP. Numbers are the summation across three separate transformations. (C) Five-fold serial dilution spot assays of indicated strains with and without 64 μg/mL fluconazole (FLZ). Two independent clones are shown for erg3Δ (BleMX). Images were captured at 48 hours.

FIG 4.

The CRISPR-RNP system efficiently generates gene deletions utilizing KanMX for C. glabrata. (A) Schematic of pUG6 plasmid. P1 and P2 indicate location of amplification primer sequences. (B) Total number of positive transformants using KanMX with and without addition of CRISPR-RNP. Numbers are the summation across three separate transformations. (C) Five-fold serial dilution spot assays of indicated strains with and without 64 μg/mL fluconazole (FLZ). Images were captured at 48 hours.

The CRISPR-Cas9 RNP system efficiently generates gene deletions utilizing the KanMX drug resistance cassette in C. glabrata.

After determining that our CRISPR-Cas9 RNP was effective for single and double gene deletions using NatMX, HphMX and BleMX, we tested whether this system permitted the use of KanMX as a drug resistance cassette in C. glabrata. Although KanMX is routinely used in S. cerevisiae, KanMX has not been successfully utilized for genetic manipulations in C. glabrata. The use of KanMX drug resistance cassettes would allow the repurposing of many S. cerevisiae tagging and deletion KanMX cassettes for C. glabrata. In our studies, we have successfully generated several C. glabrata deletion strains using HphMX or NatMX using ~130–150bp homology with chemical transformation and electroporation (36, 37). However, any attempts to use KanMX as a drug selection cassette using these two methods were not successful (data not shown and Fig. 4B). To first determine whether the CRISPR-Cas9 RNP system is sufficient for repurposing KanMX for use in C. glabrata, ADE2 was deleted using the KanMX drug resistance cassette amplified from the pUG6 plasmid (Fig. 4A) (38). We observed 55% red colonies suggesting that the CRISPR-Cas9 RNP method is efficient in generating gene deletions using KanMX and 800 μg/ml G418 (Fig. 4B). In contrast, when using electroporation without the aid of CRISPR-Cas9, only one red colony out of 86 was observed indicating an efficiency of 1.1%. We were also able to successfully delete ERG3 with our CRISPR-Cas9 RNP method using KanMX. A five-fold serial dilution spot assay with and without 64 μg/mL fluconazole were performed to compare the phenotype to previously constructed erg3Δ strains. Importantly, we observe an azole resistant phenotype similar to the other constructed erg3Δ strains (Fig. 4C). These data demonstrate that KanMX is an effective drug resistance cassette for use in C. glabrata when using the CRISPR-Cas9 RNP approach.

The CRISPR-RNP system generates endogenous epitope tagged proteins using KanMX in C. glabrata.

Because our data indicate that KanMX is a suitable drug resistance cassette for gene deletions in C. glabrata, we next determined if using the CRISPR-Cas9 RNP method would also permit endogenous epitope tagging using the C-terminal 3xHA-KanMX plasmid (pFA6a) commonly used for S. cerevisiae (Fig. 5A) (39). ERG3 and ERG11 were used to demonstrate that the CRISPR-Cas9 RNP method would allow for endogenous C-terminal tagging using KanMX. After confirming the presence of the insert via colony PCR, strains were grown with and without 64 μg/mL fluconazole in SC media and collected at mid-log phase for immunoblotting using anti-HA (12CA5). Histone H3 was used as a loading control. Our data indicate that Erg3 protein is expressed under untreated conditions and induced under fluconazole treatment (Fig. 5B), which is consistent with transcript analysis from our previous study (40). Erg11 protein is also expressed under untreated conditions and induced under fluconazole treatment (Fig. 5C) similar to what is observed for Erg3 (Fig. 5B) and consistent with previous transcript and protein analysis (40, 41). To confirm that the epitope tag does not alter azole susceptibility, we performed five-fold serial dilution spot assays with and without 64 μg/mL fluconazole, using an erg3Δ strain as a control. All epitope tagged Erg3-3xHA strains grow similar to WT under both untreated and fluconazole treatment (Fig. 5D). We observe the same effect for Erg11-3xHA strains with and without fluconazole treatment (Fig. 5E). Altogether, these data suggest that the CRISPR-Cas9 RNP system effectively generates endogenous epitope tagged proteins using KanMX and C-terminally tagging Erg11 and Erg3 does not appear to alter azole susceptibility.

FIG 5.

The CRISPR-Cas9 RNP system generates endogenous epitope tagged proteins using KanMX in C. glabrata. (A) Schematic of pFA6-3HA-KanMX plasmid. P1 and P2 indicate location of amplification primer sequences. (B and C) Indicated strains were either untreated (−) or treated (+) with 64 μg/mL of fluconazole (FLZ) for three hours. Whole cell extracts were isolated and immunoblotted against HA antibody for detection of Erg3 or Erg11. Histone H3 was used as a loading control. Three independent clones were represented for Erg3-3xHA and Erg11-3xHA. (D and E) Five-fold serial dilution spot assays of indicated strains with 0, 16, and 64 μg/mL fluconazole (FLZ), respectively. Three independent clones were represented for Erg3-3xHA and Erg11-3xHA. Images were captured at 48 hours.

The CRISPR-Cas9 RNP system allows the use of BleMX as a drug resistance cassette for C. auris.

It has been previously demonstrated that a CRISPR-Cas9 RNP system can be utilized in C. auris using SAT1 as a drug resistance cassette (14–16, 18). With this, we tested whether the CRISPR-Cas9 RNP system allowed for the utilization of BleMX as a drug resistance cassette in C. auris. We first codon-optimized BleMX for use in CTG-clade species and named the plasmid pCdOpt-BMX (Fig. 6A). Using this codon-optimized BleMX plasmid as a template, we deleted ERG3 in C. auris AR0387 using the CRISPR-Cas9 RNP method. After confirming the presence of BleMX via colony PCR, we performed five-fold serial dilution spot assays with and without 64 μg/mL fluconazole on each strain. Similar to the C. glabrata erg3Δ strains, we observed a similar azole resistant phenotype across all clones (Fig. 6B). These data determine for the first time the effective use of our codon optimized BleMX in C. auris when using the CRISPR-RNP approach.

FIG 6.

The CRISPR-Cas9 RNP system generates gene deletions using a codon optimized BleMX in C. auris. (A) Schematic of pCdOpt-BMX plasmid. P1 and P2 indicate location of amplification primer sequences. (B) Five-fold serial dilution spot assays of indicated C. auris strains with and without 64 μg/mL fluconazole (FLZ). Four independent clones were represented for Caurerg3Δ strain (BleMX). Images were captured at 48 hours.

Using CRISPR-Cas9 RNP system to generate heterozygous and homozygous deletions in C. albicans.

SET1, a known histone methyltransferase, when deleted in C. glabrata or S. cerevisiae alters azole susceptibility and ERG gene expression including ERG3 (37, 42). We sought to determine whether loss of SET1 in C. albicans exhibits a similar phenotype. To test this, we generated both heterozygous and homozygous SET1 deletion mutants in a sequential manner where the entire open reading frame was deleted with the SAT1 selection marker to generate the heterozygous deletion and then subsequently recycled using FLP recombinase to make the homozygous deletion in the C. albicans SC5314 strain (Fig. 7A) (43). Because Set1 is a histone H3K4 methyltransferase, we wanted to confirm the loss of methylation in the set1Δ/Δ strains using immunoblot analysis using H3K4me1, H3K4me2, and H3K4me3 specific antibodies (Fig. 7B). Since Set1 is the sole histone H3K4 methyltransferase in most yeast species, we observed a complete loss of H3K4 methylation in the set1Δ/Δ strain which is consistent with previous reports in a CAI4 strain (44, 45). We also determined that the SET1/set1Δ strain exhibited no change in H3K4 methylation status indicating loss of one allele didn’t impact global histone methylation (Fig. 7B). To assess azole susceptibility in C. albicans, we performed five-fold serial dilution spot assays with and without 0.5 μg/mL fluconazole. Interestingly, we did not observe altered susceptibility to azoles in the set1Δ/Δ or SET1/set1Δ strains (Fig. 7C). This is in clear contrast to what is observed when SET1 is deleted in C. glabrata and S. cerevisiae indicating a species-specific difference and utilization of SET1 (36, 37).

FIG 7.

The CRISPR-Cas9 RNP system is used for deleting SET1 in C. albicans. (A) Schematic of pBSS2-SAT1-FLP plasmid. P1 and P2 indicate location of amplification primer sequences. (B) Whole cell extracts were isolated from indicated C. albicans strain SC5314 and immunoblotted against methyl-specific H3K4 mono-, di- and trimethylation antibodies. Histone H3 was used as a loading control. (C) Five-fold serial dilution spot assays of indicated C. albicans strains with and without 0.5 μg/mL fluconazole (FLZ). Images were captured at 24 hours.

DISCUSSION

In this study, we have expanded the toolkit in Candida by utilizing the CRIPSR-Cas9 RNP approach which allowed the repurposing of drug resistance cassettes for genetic manipulation in prototrophic strains. Using these tools, we established that deleting ERG3 in C. glabrata and C. auris confers a fluconazole resistant phenotype. We also identified a synthetic genetic interaction between C. glabrata ERG3 and ERG5 and determined azole susceptible differences between C. albicans set1Δ/Δ strains and C. glabrata set1Δ strains.

We show that the use of CRISPR-Cas9 RNP with homology regions of 130–150 bp is efficient at making single gene deletions, double deletions and epitope tags in C. glabrata. Although three CRISPR-Cas9 RNP studies have used long flanking sequence for gene deletions in C. glabrata and C. auris (14, 15, 18), studies have shown short homology regions of ~50–70bp are feasible for making gene deletions in C. auris and C. albicans (16, 17, 46). We have attempted to use short homology regions of ~60 bp for deleting genes in C. glabrata, but it appears not to be as consistent using ~130–150 bp flanking sequences. In addition, 130–150 bp homology regions have been useful for making double deletion and epitope tagged strains in C. glabrata.

Using the CRISPR-Cas9 RNP method, we also determine that two additional drug resistance cassettes (KanMX and BleMX) can be used reliably in C. glabrata allowing for more complex genetic manipulations. With a repertoire of four drug resistance cassettes available for use in C. glabrata, this greatly increases the flexibility and utility for manipulating prototrophic clinical isolates where auxotrophic makers are not readily available or feasible. In addition, our study successfully demonstrates the repurposing of KanMX-containing plasmids traditionally utilized for making gene deletions or C-terminal epitope tags in S. cerevisiae, for use in C. glabrata. While we clearly demonstrate that the endogenous C-terminal 3xHA tagging constructs used for S. cerevisiae is suitable for C. glabrata, this approach may not work for all genes, as C-terminal tagging may disrupt the function of the protein. Thus, our approach would also allow for repurposing endogenous N-terminal tagging constructs designed for S. cerevisiae. For example, our lab has generated N-ICE plasmids with KanMX selection cassettes for N-terminal tagging essential and non-essential genes in S. cerevisiae (47). We would suspect these plasmids and other KanMX-containing plasmids could be directly used in C. glabrata. Additionally, the efficiency of endogenous epitope tagging proteins using CRISPR allows for more functional and mechanistic studies, as endogenous epitope tagged proteins have been used sparingly in prototrophic strains and clinical isolates of C. glabrata. This is particularly important since antibodies to endogenous proteins are scarce and costly to make.

Our study also shows that BleMX dominant drug selection cassette can be used in deleting genes in C. glabrata but also C. auris. Although BleMX has been used previously in C. glabrata using standard electroporation, the efficiency was extremely low and has not been typically used for routine genetic manipulation (34). BleMX showed the lowest efficiency of the drug selection cassettes used in our study. Nonetheless, we clearly demonstrate the CRISPR-Cas9 RNP method does improve homologous recombination efficiency enough where BleMX can be used. Moreover, our codon optimized BleMX plasmid will be readily available as another effective and needed dominant selection cassette for C. auris. Currently, we have not determined if our codon optimized BleMX drug resistance cassette can be used in other CTG clade species.

CRISPR-Cas9 RNP has been used successfully for C. glabrata, C. auris, C. lusitaniae, and C. albicans (14–18). We have also successfully used the CRISPR-Cas9 RNP approach to delete C. albicans SET1 using a recyclable SAT1 cassette. Interestingly, the loss of SET1 does not confer azole susceptibility in contrast to when SET1 is deleted in C. glabrata or S. cerevisiae which is due to either altered ERG11 gene expression or PDR5 expression, respectively (36, 37). Because C. albicans is part of the CTG clade and is more evolutionarily distant to C. glabrata and S. cerevisiae, this may suggest a species-specific utilization of SET1.

In contrast to C. glabrata, we have not been able to utilize KanMX in C. albicans due this organism’s high tolerance/resistance to the aminoglycoside antibiotic, G418. However, it has been reported that adjuvants such as quinine or molybdate can suppress background growth of C. albicans and allow successful integration of codon optimized CaKan and CaHygB cassettes using standard chemical transformation procedures (48). We anticipate that using these constructs, adjuvants, and the CRISPR-Cas9 RNP method could reduce background growth and increase HDR for efficient use of these markers in C. albicans. Alternatively, simultaneous deletion of both alleles with KanMX or HygB without adjuvants may work, since a CRISPR-RNP based system has been used successfully to simultaneously delete both alleles in C. albicans when using SAT1 and HygB (17).

Overall, our study provides the field additional ways to efficiently manipulate Candida pathogens. Importantly, this approach provides us further insight in the ergosterol pathway and species differences in azole susceptibility in Candida pathogens when SET1 is deleted although additional studies would be needed to further address the mechanisms of these observations. Applying this expanded toolkit to other studies in Candida should enhance our understanding of fungal drug resistance and pathogenesis.

MATERIALS AND METHODS

Yeast strains and plasmids

All strains used are described in Table S1. C. glabrata strains were derived from the Cg2001 (ATCC 2001). C. albicans strains were derived from SC5314 (49), a gift from William A. Fonzi, Georgetown University. C. auris AR0387 strain was obtained from the CDC AR Isolate Bank. Yeast cells were grown in YPD medium or synthetic complete (SC, Sunrise Science) medium as indicated. The pAG25, pAG32, and pUG6 plasmids were obtained from Euroscarf (22, 38). The pFA6a-3HA-KanMX and pCY3090-07 plasmids were obtained from Addgene (35, 39). The pBSS2-SAT1 flipper plasmid was provided to us by P. David Rogers, St. Jude Children’s Research Hospital with permission from Joachim Morschauser (43). pCdOpt-BMX (BleMX) was synthesized by IDT where the TEF1p-BleMX-TEF1t sequence was codon optimized for CTG clade Candida species using the IDT codon optimization tool, custom synthesized and cloned into the pUCIDT plasmid. The pCdOpt-BMX plasmid can be obtained at Addgene (ID number 203929).

PCR amplification for gene deletion and epitope tagging

All oligonucleotides used are denoted in Table S2. Forward primers used for gene deletions were designed with homology regions of ~130–150 bp flanking the 5’-ORF of the target gene of interest followed by 20–25 base pairs of sequence homologous to the indicated plasmid. Reverse primers were designed with homology regions ~130–150 bp flanking the 3’-ORF of the target gene of interest followed by 20–25 base pairs of sequence homology to the indicated plasmid. PCR conditions for amplification of replacement cassettes are as follows: 95°C for 5 minutes; 95°C for 30 seconds, 52°C for 30 seconds, 72°C for 2–3 minutes for a total of 30 cycles, with a final elongation step at 72°C for 10 minutes. The final PCR products were pooled and purified from agarose gels.

CRISPR gRNA design and selection

Custom Alt-R CRISPR gRNAs were designed and ordered from Integrated DNA Technologies (Table S3). For each gene deletions, two CRISPR gRNAs were designed in close to the 5’ and 3’ ORF of the gene of interest. For epitope tagging, one CRISPR gRNA was designed in the 3’UTR of the gene of interest. CRISPR gRNAs were selected based upon their designated “On-Target Score” as determined by the CRISPR-Cas9 guide RNA design checker (IDT). Potential gRNAs were screened for Off-Target events using the CRISPR RGEN Tools Cas OFFinder (http://www.rgenome.net/cas-offinder/). Selected gRNAs required >75 On-Target Score as well as 0 potential off target events with 3 mismatches or less.

CRISPR-Cas9 RNP system

The CRISPR-Cas9 RNP method was based on Grahl et al. with slight modifications (18). Briefly, Alt-R CRISPR crRNA and tracrRNA were used at a working concentration of 20 μM. CRISPR-Cas9 crRNAs:tracrRNA hybrid was made by mixing together 1.6 μL of crRNA (8 μM final concentration), 1.6 μL of tracrRNA (8 μM final concentration), and 0.8 μL of RNAse free water. Two crRNAs, 0.8 μL of each was added at a stoichiometric equivalent to tracrRNA and for C-terminal tagging one crRNA, 1.6 μL was used. The CRISPR RNP mix was incubated at 95°C for 5 minutes and allowed to cool to room temperature. 3 μL of 4 μM Cas9 (IDT) was added to the mix (final concentration of 1.7 μM) and incubated at room temperature for 5 minutes.

Cell transformation

25 mL of the desired strain was grown to an OD₆₀₀ of 1.6 to saturation prior to transformation. Cells were collected by centrifugation, resuspended in 10 mL 1x LiTE Buffer (100 mM LiAc, 10 mM Tris-HCl, 1 mM EDTA), and shaken at 250 rpm at 30°C for an hour. DTT was added to a final concentration of 100 mM and cells were incubated at 30°C for an additional 30 minutes. Cells were then collected by centrifugation, washed twice with 1 mL ice cold water, and washed once more with 1 mL of cold sorbitol. Cells were resuspended in 200 μL of cold sorbitol for electroporation.

Electroporation and colony PCR

20 μL of prepared cells, 1–3 μg of drug resistant cassette DNA, CRISPR mix, and RNAse free water to a final volume of 45 μL was mixed and transferred to a BioRad Gene Pulser cuvette (0.2 cm gap). Cells were pulsed using an Eppendorf Eporator at 1500 V and immediately resuspended in 1 mL of ice-cold Sorbitol. Cells were then collected by centrifugation, resuspended in 1 mL of YPD media, and allowed to recover by incubation at 30°C at 250 rpm for 3–24 hours. Cells were then collected, resuspended in 100 μL of YPD, and plated onto drug selective media at the desired concentration. Nourseothricin (GoldBio) was used at a final concentration of 300 μg/mL for antibiotic selection of the NatMX cassette. Hygromycin B (Cayman) was used at a final concentration of 500 μg/mL antibiotic selection of the HphMX cassette. Geneticin (G418, GoldBio) was used at a final concentration of 800 μg/mL for antibiotic selection of the KanMX cassette. Zeocin (Cayman) was used at a final concentration of 600 μg/mL for antibiotic selection of the BleMX cassette in C. glabrata and 800 μg/mL in C. auris. Colonies were streaked onto fresh plates with the desired drug, and single colonies were selected and restreaked onto fresh YPD plates. Colonies were screened via colony PCR using primers indicated in Table S2. Three independent clones were verified by PCR and analyzed for phenotypic characterizations.

Serial dilution growth assay

For serial dilution spot assays, yeast strains were inoculated in SC media and grown to saturation overnight. Yeast strains were diluted to an OD₆₀₀ of 0.1 and grown in SC media to log phase shaking at 30°C. The indicated strains were spotted in five-fold dilutions starting at an OD₆₀₀ of 0.01 on untreated SC plates or plates containing 8 μg/mL, 16 μg/mL, or 64 μg/mL fluconazole (Cayman). For C. glabrata and C. auris, plates were grown at 30°C on SC plates for 48 hours prior to imaging. For C. albicans, plates were grown at 30°C on YPD plates for 48 hours prior to imaging.

Quantitative real-time PCR analysis

RNA was isolated from cells grown in SC media by standard acid phenol purification as previously described (37). Reverse transcription to generate cDNA was performed using the ABM All-In-One 5X RT MasterMix (ABM). Primer Express 3.0 software was used for designing primers for gene expression analysis by quantitative real-time polymerase chain reaction (Table S4). A minimum of three biological replicates, as well as three technical replicates, were performed for each biological replicate. All data were analyzed using the comparative CT method (2^−ΔΔCT). RDN18 (18S ribosomal RNA) was used as an internal control. All samples were normalized to the Cg2001 WT strain (Table S5).

Cell extract and Western blot analysis

Whole cell extraction and Western blot analysis were performed as previously described (50, 51). The anti-HA (Roche 12CA5, 1:10,000) monoclonal antibody was used as previously described (52). Histone H3K4 methylation-specific antibodies were used as previously described: H3K4me1 (Upstate, 07-436; 1:2,500), H3K4me2 (Upstate, 07-030; 1:10,000), and H3K4me3 (Active Motif 39159, 1:100,000) (42, 53). Histone H3 rabbit polyclonal antibody (PRF&L) was used at a 1:100,000 dilution.

IMPORTANCE:

The increasing problem of drug resistance and emerging pathogens is an urgent global health problem that necessitates the development and expansion of tools for studying fungal drug resistance and pathogenesis. We have demonstrated the effectiveness of an expression-free CRISPR-Cas9 RNP-based approach employing 130–150 bp homology regions for directed repair. Our approach is robust and efficient for making gene deletions in C. glabrata, C. auris and C. albicans as well as epitope tagging in C. glabrata. Furthermore, we demonstrated that KanMX and BleMX drug resistance cassettes can be repurposed in C. glabrata and BleMX in C. auris. Overall, we have expanded the toolkit for genetic manipulation and discovery in fungal pathogens.