Yeast, monocellular eukaryotes, are easy to culture and grow rapidly. Compared with other eukaryotes, yeast have a simpler genetic background. Yeast are also used in industrial fermentation owing to their easy operation and safety. CRISPR technology has been widely used in yeast for the following main objectives: 1) yeast polygene and polyploid knockout, 2) the synthesis of yeast chromosomes, 3) genome-scale engineering and library creation, 4) gene integration, heterologous expression, and metabolic pathway regulation, 5) CRISPR-mediated activation/interactions, 6) non-traditional gene editing and explorations of virulence mechanisms.
2.1.1 Polygene and polyploid editing of yeast
Rapid advances in gene sequencing technology have resulted in substantial improvements in our understanding of microbial genomes. The existence and distribution of functional genes can be deciphered by the gene-editing method, providing a basis for in-depth studies of fungal microbiome functions (Aguilar-Pontes et al. 2014). Traditional yeast polygenic editing requires multiple rounds of selective pressure (Adames et al. 2019). CRISPR technology can reduce or even avoid the use of resistance. In 2013, DiCarlo et al. (2013) first introduced the CRISPR/Cas9 system into Saccharomyces cerevisiae, demonstrating an improvement in the efficiency of double-strand break (DSB) repair by gene recombination by 130-fold. Bao et al. (2015) developed a homology-integrated CRISPR/Cas (HI-CRISPR) system in S. cerevisiae. They realized one-step multigene disruption by concatenating different guide RNAs (gRNAs) on a vector, with an efficiency of 27–87%. Mans et al. (2015) transformed three plasmids by the in vitro assembly of plasmids containing two gRNAs, and simultaneously produced six gene deletion strains of S. cerevisiae. The CRISPR/Cas system has also been used for the multiple genome engineering of polyploid industrial yeasts. Zhang et al. (2014) used Cas9 to knock out four genes of S. cerevisiae ATCC 4124 with 15–60% efficiency, and four nutrient-deficient strains were obtained. Lian et al. (2017) expressed gRNA by high-copy plasmids and then transformed the plasmids into a diploid strain (Ethanol Red) and a triploid line (ATCC 4124). Four genes were knocked out in a single step with an efficiency of 100%. Additionally, Li et al. (2018) found that Cpf1 from Francisella novivida (FnCpf1) can target DNA fragments assembled in vivo for singleplex, doubleplex, and tripleplex genomic integration, with efficiencies of 95%, 52%, and 43%, respectively. Yang et al. (2020) used CRISPR/Cpf1 to target the CAN1 and URA3 genes of Yarrowia lipolytica, obtaining efficiencies of up to 93% and 96%. Zhao et al. (2020) also applied the CRISPR/Cas12a system to Schizosaccharomyces pombe. Because Cpf1 does not need tracrRNA and lacks the HNH endonuclease domain present in Cas9, it is well-suited to yeast multiplex gene editing (Zetsche et al. 2015; Swiat et al. 2017; Zetsche et al. 2017; Li et al. 2018). Multi-gene editing technology based on CRISPR can be used to create complex traits and explore gene regulatory networks.
2.1.2 Synthesis of yeast chromosomes
Variation in chromosome structure plays an important role in the diversity of biological characters. The synthetic yeast genome, designated Sc2.0, was an international cooperative project aimed at designing and constructing the first fully chemically synthesized eukaryotic cell with 16 chromosomes (Richardson et al. 2017). CRISPR/Cas9 provides a method for the redesign and construction of S. cerevisiae chromosomes. Chromosome drives via CRISPR/Cas9 enable the biased inheritance of complex genetic traits on a chromosomal scale in yeast (Xu et al. 2020). Luo et al. (2018) successfully fused yeast chromosomes using CRISPR/Cas9 and produced a series of homologous strains with 2 to 16 chromosomes. Xie et al. (2017) constructed a ring synthetic chromosome Ⅴ of S. cerevisiae by CRISPR technology. Shao et al. (2018) used CRISPR/Cas9 to efficiently knock out redundant centromeres and telomeres of chromosomes for fusion in S. cerevisiae and used homologous recombination to realize 15 rounds of chromosome fusion. Finally, 16 natural chromosomes of a haploid S. cerevisiae were artificially fused into the yeast strain SY14 with a single chromosome. In 2019, the CRISPR/Cas9 method was further used to induce DSBs in the proximal regions of two telomeres of the SY14 linear chromosome, and the ends of two DSBs were connected with donor DNA fragments by endogenous homologous recombination to generate a new strain with a single circular chromosome, designated SY15 (Shao et al. 2019). The rearrangement of yeast chromosomes clearly establishes the effectiveness of CRISPR/Cas-based systems for evaluating the structure and function of genes in eukaryotic evolution.
2.1.3 Genome-scale engineering of yeast and library creation
The programmable control of gene expression is essential for understanding gene function, regulating cell behavior, and developing therapeutic methods. Genetic screening can be used to study the functions of multiple genes simultaneously in a high-throughput manner. CRISPR/Cas9 has been used as a screening strategy to induce mutations and assess gene function. In 2018, Bao et al. (2018) developed CRISPR/Cas9 and homology-directed-repair assisted genome-scale engineering (CHAnGE), which can be used to edit the whole genome of S. cerevisiae at the single nucleotide level. This method can rapidly generate thousands of specific mutant yeasts. CHAnGE can generate single base pair changes on the whole chromosome and minimize the impact on the function of adjacent genes with unprecedented accuracy. The authors have also established a gene knockout yeast library using the system to improve the production of heterologous natural products. Li et al. (2019) created the CRISPR Activation Library based on the gene regulation system of dCas9. They identified a high-temperature resistant yeast with a new gene regulatory mechanism by the construction of a library with 260 sgRNAs. Guo et al. (2018) used CRISPR/Cas9 to induce DSBs in yeast target genes and provided a donor template with programmed mutations for homology-directed repair (HDR), realizing the high-throughput creation and functional analysis of a yeast DNA sequence variant library. Ferreira et al. (2019) used dCas9-VPR and a library of 3194 gRNAs targeting 168 genes to screen targets for enhancing the flux toward cytosolic malonyl-CoA, significantly increasing the production of 3-hydroxypropionic acid (3-HP). Buchmuller et al. (2019) constructed a yeast library by CRISPR/Cpf1-assisted tag library engineering (CASTLING). The yeast library is an ideal model for studies in systematic biology, and the CRISPR/Cas system is a practical tool for the establishment of systematic libraries and for genome-scale research.
2.1.4 Gene integration, heterologous expression, and metabolic pathway regulation
Balancing gene expression levels in endogenous or exogenous metabolic pathways is an important way to improve the yield and efficiency of target products in yeast (Idiris et al. 2010). Eukaryotes can produce abundant secondary metabolites (Jiang et al. 2021; Liu et al. 2021; Sharma et al. 2021). There secondary metabolic clusters are typically silent and transcriptionally inactive under laboratory conditions(Osbourn 2010; Brakhage and Schroeckh 2011; Keller 2019). These silent gene clusters have been cloned and transferred into S. cerevisiae or other microorganisms to activate the expression of gene clusters (Keller 2019). The target gene was cleaved by a nuclease to form DSBs and then repaired by HDR or non- homologous end joining (NHEJ) (Ceccaldi et al. 2016). NHEJ is feasible for the construction of mutant libraries, since repair is random. However, for rational metabolic engineering, HR is preferred owing to its precision and predictability. CRISPR/Cas enables precise gene editing, which can undoubtedly accelerate the heterologous expression of silent genes (Jakounas et al. 2015; Xu et al. 2016). Both HR and NHEJ activity via CRISPR/Cas9-induced oligodeoxynucleotide (ODN)-mediated DSB repair have been quantitatively measured (Du J et al. 2018). Horwitz et al. (2015) used the CRISPR system to integrate six DNA fragments carrying 11 genes with a total length of 24 kb in S. cerevisiae and finally established prototype pathways for muconic acid production. In 2016, Jessop-Fabre et al. (2016) constructed the EasyClone-MarkerFree plasmid toolbox, which can simultaneously insert 1–3 DNA fragments into the genome of S. cerevisiae without using selective markers. Ronda et al. (2015) combined the stability and versatility of the EasyClone vector system with the precision and efficiency of CRISPR/Cas9 to efficiently integrate three genes involved in the β-carotene pathway at three different sites on three chromosomes in S. cerevisiae, thus improving the production of β-carotene. Shi et al. (2015) constructed a Delta integration CRISPR/Cas (Di-CRISPR) platform for multi-copy gene integration, which can efficiently integrate large-scale biochemical pathways without labeling. Utilizing this method, they achieved the unprecedented one-step integration of 18-copy genomes of 24 kb and successfully constructed a yeast strain capable of directly using xylose to produce (R,R)-2,3-butanediol (BDO). Wan et al. (2018) overexpressed MRP8, which encodes a mitochondrial ribosomal protein, in recombinant S. cerevisiae Y294 by CRISPR/Cas9 technology, thereby increasing extracellular Cbh1 enzyme activity by 80%. Sun et al. (2020) successfully knocked out two copies of the gene encoding isocitrate dehydrogenase (ICD) in Pichia kudriavzevii and integrated At_CAD and Pk_MTTA in the mutant strain to increase the yield of itaconic acid (IA). CRISPR/Cas9 technology has also been used to remove genes with adverse effects. For example, Chin et al. (2016) mutated the yeast CAR1 gene, encoding arginase, by CRISPR/Cas9 technology, thus reducing carcinogenic ethyl carbamate (EC) produced during ethanol fermentation.
Due to differences in the recognition of promoters among eukaryotic taxa, silent gene clusters cannot be activated when they are directly cloned and transferred into heterologous hosts. The CRISPR/Cas9 system can be used to change the original regulatory elements to achieve the heterologous expression of these genes. Kang et al. (2016) described an improved yeast-based promoter engineering platform (mCRISTAR) that combines CRISPR/Cas9 and the transformation-associated recombination (TAR) (Yamanaka et al. 2014) to replace natural promoters with combinatorial promoters. CRISPR/Cas9 mediates DSBs to form linear DNA fragments at the target promoter and then integrates the homologous arm with biosynthetic gene clusters (BGCs) by homologous recombination in yeast cells. Up to 32 promoters can be inserted into a single natural BGC by four rounds of mCRISTAR using four auxotrophic markers commonly used in yeast, and silent gene clusters can be transcriptionally activated (Kang et al. 2016).
2.1.5 CRISPR activation (CRISPRa)/interaction (CRISPRi) in yeast
Recently, D10A and H804A mutations were introduced into the Ruvc and HNH domains of the Cas9 protein in the CRISPR system to obtain nuclease-deficient Cas9 (dCas9). By fusing dCas9 with different types of transcriptional regulatory domains, such as transcriptional inhibitors, activators, or epigenetic modification enzymes, dCas9 proteins can target specific sites of target genes, resulting in different modes of gene regulation (Larson et al. 2013; Qi et al. 2013) (Fig. 2). Changes in gene expression and activity can be obtained by CRISPRa/CRISPRi in yeast. dCas9-Mxi1 and dCas9-VPR are the most commonly used regulators (Jensen 2018). Gilbert et al. (2013) fused dCas9 with Mxi1, a mammalian transcription inhibition domain, and targeted the Tef1 promoter. They found that the inhibitory effect of dCas9 on reporter gene expression increased from 18-fold for dCas9 alone to 53-fold, indicating that dCas9 binding to transcriptional regulators is highly effective. Schwartz et al. (2017) applied the CRISPRi system to Yarrowia lipolytica, designed sgRNAs for the TSS and TATA box of target gene promoter regions, and successfully inhibited 8 of 9 target genes. They also found that dCas9-mxi1 increased the inhibition of KU80 gene transcription from 38% for dCas9 to 87%. Vanegas et al. (2017) combined Cas9 and dCas9 into a SWITCH dynamic CRISPR tool where switching mechanism is based on the recombination of dCas9 after Cas9 is directed to cleave its own gene sequence. The tool enables S. cerevisiae strains to alternate between genetic engineering and metabolic pathway control states, enabling the accurate control of multiple genes of S. cerevisiae.
Farzadfard et al. (2013) fused dCas9 with VP64 (a commonly used eukaryotic transcription activator domain) in S. cerevisiae and targeted to a minimal CYC1 promoter (pCYC1m). The expression of gRNAs in different regions of pCYC1m resulted in different levels of fluorescent GFP reporter gene activation and inhibition, indicating that dCas9/dCas9-VP64 can interfere with the formation of the transcription initiation complex by targeting different positions in endogenous promoters. Lian et al. (2017) tested a combination of dCas proteins from different sources (S. pyogenes, Staphylococcus aureus, Streptococcus thermophiles, and Lachnospiraceae bacterium) , and multiple transcriptional activation or inhibitory domains and finally constructed a three-phase gene regulatory strategy for S. cerevisiae based on dLbCpf1-VP (CRISPRa), dSpCas9-RD1152 (CRISPRi), and SaCas9 (CRISPRd), known as CRISPR-AID. Using this system, they transformed a single plasmid into yeast to simultaneously induce a 5-fold increase in red fluorescent protein, 5-fold inhibition of yellow fluorescent protein, and 95% deletion of the endogenous gene. These studies demonstrate that CRISPR/Cas is a powerful tool for fungal genetic screening. CRISPRa/i reversibly regulates the expression of target genes rather than mutation, which reduces errors in gene repair.
2.1.6 Development of a CRISPR gene editing platform in non-traditional yeast and exploration of virulence mechanisms
The successful application of CRISPR/Cas9 in S. cerevisiae genome editing has prompted interest in the genetic manipulation of various non-traditional yeasts (Arras et al. 2016; Schwartz et al. 2016; Raschmanová et al. 2018). In particular, the CRISPR/Cas9 system has been developed to study the virulence mechanisms of Candida albicans, Candida glabrata, Candida parapsilosis, Cryptococcus neoformans, and other clinically related pathogenic yeasts (Vyas et al. 2015; Enkler et al. 2016; Lombardi et al. 2017; Wang 2018). In 2015, Vyas et al. (2015) first applied the CRISPR/Cas9 system to gene editing in C. albicans. By introducing the Cas9 protein and sgRNA, the target gene was driven by RNA polymerase III promoter SNR52, and a repair template containing a stop codon was provided, resulting in the production of homozygous mutants in one transformation. Xu et al. (2018) successfully obtained a homozygous inactivation mutant of CaMIT1 by the CRISPR/Cas9 method. The mutant strain was sensitive to calcium and lithium ions, sodium dodecyl sulfate, clotrimazole, and ketoconazole but tolerant of Congo red. Min et al. (2016) showed that CRISPR/Cas9 gene elements can play a transient role in C. albicans without stable integration into the genome, addressing the concern that Cas9 may cause long-term adverse effects (such as off-target effects) in the C. albicans genome. Huang et al. (2017) developed a marker recovery strategy using CRISPR/Cas9. Two marker genes can be used to sequentially screen homozygous deletion mutants with three or more genes in the same strain. Nguyen et al. (2017) removed CRISPR and nourseothricin (NAT) markers from the genome by the SAT1 flipper system after determining the target site modification of C. albicans, thus allowing the next round of unlabeled genome editing. Grahl et al. (2017) used a purified Cas9 protein, crRNA, and tracrRNA to form ribonucleoproteins (RNPs) to modify the genome of three non-C. albicans Candida (NCAC), resolving the low promoter activity of the CRISPR/Cas9 system for different species-specific patterns of gene expression. Rybak et al. (2020) introduced TAC1B mutations from drug-resistant clinical isolates into the fluconazole-susceptible C. auris strains by an RNP-mediated transformation system and found that TAC1B mutations explain the observed increase in fluconazole resistance. Lombardi et al. (2017) described the first CRISPR/Cas9 editing system based on plasmids with an autonomous replication sequence (ARS) in C. parapsilosis. The gRNA is released between two ribozymes (Hammerhead and hepatitis delta virus [HDV]) for multiple editing of the target gene. In 2019, they improved the system so that the gRNA could be introduced in a single cloning step and released by cleavage between a tRNA and a ribozyme (Lombardi et al. 2019). This method was used for efficient gene editing in C. parapsilosis, C. orthopsilosis, and C. metapsilosis. Zhang et al. (2019) developed a CRISPR/Cas9 system for genome editing in C. tropicalis, demonstrating the efficient deletion of single or double genes in 9 days, 17 fewer days than required for the traditional SAT1 flipper strategy. This method can be used to promote the assembly and stable integration of multiple DNA fragments into a target site in C. tropicalis. Zoppo et al. (2020) used CRISPR/Cas9 technology to study the virulence of Als in C. parapsilosis. Ibrahim et al. (2020) constructed an episomal vector for the expression of Cas9 and sgRNA by using an ARS isolated from C. aaseri SH14 and used a single sgRNA with 70% efficiency to destroy six copies of acyl-CoA oxidase genes (AOX2, AOX4, and AOX5) in diploid cells simultaneously.
2.2 Filamentous fungi
Filamentous fungi play important roles in the biomass cycle of ecosystems. They can produce a variety of secondary metabolites, such as cellulase, pectinase, and protease (Chandel et al. 2012; Li et al. 2020). These fungi can result in the deterioration of buildings, food, and feed and can even cause fatal diseases in humans (Nevalainen et al. 2015; Köhler et al. 2017; Avery et al. 2019). Before the CRISPR/Cas system was proposed, gene editing of these fungi was a big issue in biology (Wang et al. 2017). The CRISPR/Cas9 system has provided a convenient tool for studies of filamentous fungi, such as analyses of gene function, pathogenic mechanisms, and metabolic pathways and the development of methods to increase product yields.
2.2.1 Unlocking gene function
CRISPR/Cas9 technology has been used for genome editing in various filamentous fungi (Nødvig et al. 2015; Wang and Coleman 2019), such as Trichoderma reesei (Liu et al. 2015), Neurospora crassa (Matsu-Ura T 2015), Aspergillus oryzae (Katayama et al. 2015), Aspergillus niger (Kuivanen et al. 2016), Magnaporthe oryzae (Foster et al. 2018), Myceliophthora thermophila (Liu et al. 2017), Aspergillus nidulans (Zhang et al. 2016), Ustilago maydis (Schuster et al. 2016), Mucor circulalloides (Nagy et al. 2017), Phytophthora sojae (Miao et al. 2020), Aspergillus fumigatus (Fuller et al. 2015), and Penicillium chrysogenum (Pohl et al. 2016). The Cas9-gRNA complex assembled in vitro or Cas9 and gRNA expressed in vivo have been employed to deactivate genes, followed by comparative analyses of metabolites, cell morphology, toxicity, and other properties between the wild-type and deletion strains to verify gene functions .
Liu et al. (2015) edited T. reesei genes by optimizing the transcription of the Cas9 protein and in vitro gRNA transcription and successfully applied the CRISPR/Cas9 system to filamentous fungi for the first time in 2015. Nielsen et al. (2017) discovered a new gene in Talaromyces atroroseus responsible for the production of polyketide-nonribosomal peptide hybrid products using CRISPR/Cas9 technology. Subsequently, the application of the CRISPR/Cas9 system in fungi has been continuously optimized. Zhang et al. (2016) used the CRISPR/Cas9 system to accurately integrate the GFP gene into predicted sites in clinical isolates of A. fumigatus without inserting markers by microhomology-mediated end joining (MMEJ), and the integration efficiency was 95–100% using only a 35 bp homologous arm.
Nødvig et al. (2015) constructed a general vector and used Tef1 and GpdA promoters to drive the expression of Cas9 and gRNA, respectively. By adding nuclease sequences HH and HDV at both ends of the gRNA, gene knockout was finally achieved in six strains of fungi, including A. nidulans and A. niger. Liu et al. (2017) used the CRISPR/Cas9 system for multiple gene editing in M. thermophila. The recombination efficiency of single-gene mutations was 90–95%, that of two gene mutations was 61–70%, and those of three and four gene mutations were 30% and 22%, respectively. Zheng et al. (2018) used endogenous 5S rRNA of A. niger to drive sgRNA transcription and used 40 bp homologous donor DNA to directly delete a 48 kb long DNA fragment from the A. niger genome for the first time, with a targeting efficiency of 100%.
Owing to the limited number of selective marker genes in filamentous fungi and the difficulty in multiple rounds of gene manipulation, a single screening marker recycling method in filamentous fungi using the CRISPR/Cas9 system has been established. The self-replicating cas9 plasmid in U. maydis and P. chrysogenum would be lost in the absence of resistance pressure, thus avoiding its influence on the growth of the strain (Pohl et al. 2016; Schuster et al. 2016). Furthermore, CRISPR/Cas9 has been combined with Cre/loxP to develop a marker-free fungal gene editing system. First, the CRISPR/Cas9 system is used to break a target gene, and the repair template containing the screening marker was integrated into the cleavage site; then, Cre recombinase activated by light illumination could simultaneously delete the selective marker and CRE (Zhang et al. 2016). Liu et al. (2019) developed a V-type CRISPR/Cas12a (AsCpf1) system in M. thermophila. Through three rounds of transformation with two selectable markers, nine genes involved in the cellulase production pathway were targeted. The protein productivity and lignocellulase activity of a mutant (referred to as M9) were 9.0- and 18.5-fold higher than those of the wild type. Cas12a was also used for gene editing in A. nidulans and A. niger (Vanegas et al. 2019).
2.2.2 Interference with metabolic pathways to obtain secondary metabolites and increase yield
Some secondary metabolites of microorganisms are important pathogenic factors contributing to human and plant diseases; however, some are also important sources of bioactive substances and drug precursor compounds (Evidente et al. 2014). Filamentous fungi have a strong metabolic capacity and can produce secondary metabolites with diverse structures, such as biocidal agents, drug precursors, and antitumor bioactive substances (Hoffmeister and Keller 2007; Osbourn 2010; Ma et al. 2016). The CRISPR/Cas system can accurately alter gene expression for functional analyses of filamentous fungal metabolic gene clusters, analyses of synthesis and regulatory mechanisms, and the activation of silent gene clusters to interfere with metabolic bypass, which is expected to improve the production and activity of secondary metabolites of filamentous fungi. Kuivanen et al. (2016) combined a transcriptomics approach and CRISPR/Cas technology to delete genes involved in galactaric acid catabolism in A. niger and then heterologously expressed uronate dehydrogenase, yielding a mutant able to convert pectin-rich biomass to galactaric acid in a consolidated bioprocess.
In addition to the conventional gene editing, CRISPR/Cas technology can also knock-in strong promoters or replace promoters upstream of target genes, thus activating silent gene clusters, regulating biosynthetic genes, and synthesizing corresponding metabolites. Matsu-ura et al. (2015) successfully replaced the endogenous promoter of the cellulase related gene CLR-2 with a β-tubulin promoter in N. crassa using the CRISPR/Cas9 system. The mRNA expression of CLR-2 in the mutant strain increased about 200-fold and cellulase production increased significantly. Therefore, the CRISPR/Cas system is a powerful synthetic biology tool for the control of the biosynthesis of secondary metabolites in fungi.
2.3 Macrofungi and other fungi
Macrofungi are multicellular eukaryotes; most are binucleate or multinucleated cells. The low efficiency of obtaining homozygous mutations makes it difficult to achieve the specific modification of target genes and multigene knockout or gene insertion mutations, thereby limiting basic research focused on macrofungi (Alberti et al. 2020). At present, research on CRISPR/Cas9 in macrofungi is mainly focused on the establishment of the system in different species. In 2016, Waltz et al. (2016) used CRISPR/Cas9 to knock out one of six polyphenol oxidase (PPO) genes in Agaricus bisporus, reducing enzyme activity by 30% and effectively slowing the rate of browning. Qin et al. (2017) first used CRISPR/Cas9 technology to disrupt the URA3 gene of Ganoderma lucidum. The gene-editing efficiency of G. lucidum was proportional to the amount of gRNA (Qin et al. 2017). Liu et al. (2020) added an intron upstream of the Cas9 gene, increasing the CRISPR/Cas9-mediated gene disruption frequency in G. lucidum by 10.6-fold. Chen et al. (2018) first applied the CRISPR/Cas9 system to Cordyceps militaris by using an optimized Cas9 protein and synthesized gRNA in vitro. In 2017, Deng et al. (2017) used CRISPR/Cas9 technology tconfirmed that a SbaPKS gene in Shiraia sp. is involved in the biosynthesis of hypocrellin, related to virulence. Sugano et al. (2017) combined the cryopreserved protoplasts and CRISPR/Cas9 technology in Coprinopsis cinerea to disrupt the GFP gene by using the CcDed1 promoter to express Cas9. In addition, CRISPR/Cas9 technology has been applied to many other fungi, such as Colletotrichum gloeosporioides (Guo and An 2020), Fusarium oxysporum (Wang et al. 2018), Monilinia fructicola (Zhang et al. 2020), Sclerotinia sclerotiorum (Li et al. 2018), nematode-trapping fungi (Youssar et al. 2019), Beauveria bassiana (Chen et al. 2017), and Blastomyces dermatitidis (Kujoth et al. 2018).