A few species of filamentous fungi have been genetic models of choice since the 1950s due to their haploid growth stage, facile sexual cycles, abundant sporulation, rapid growth and, with time, large repertoires of mutants and molecular transformation systems. However, given the importance of fungi in medicine, agriculture, and ecosystems, considerable efforts have been invested over several decades to establish molecular transformation and targeted mutation systems for a much broader range of species. These include the Epichloë
species (family Clavicipitaceae, order Hypocreales), which are systemic, constitutive, and often seed-transmitted symbionts (endophytes) of cool-season grasses (Poaceae, subfam. Poöideae), and which are capable of producing a panoply of bioprotective alkaloids [1
]. However, features of important Epichloë
species that present special difficulties for genetic experimentation include growth rates far slower than model fungi, sparse sporulation, limited availability of selectable markers, and for many, diploid or triploid genomes and the lack of a sexual cycle [2
The development of CRISPR technologies has opened the door to facile gene inactivation, removal, or even replacement for a wide range of organisms, including filamentous fungi [5
]. Initially, the application of CRISPR in eukaryotes involved transformation or transfection with a gene construct expressing the Cas9 double-strand DNase and another construct to transcribe guide and tracrRNAs to direct the activity of Cas9 to the target sites. In Aspergillus
species, Nødvig et al. [7
] developed a system based on a single plasmid harboring a chimeric RNA guide and the cas
9 gene under fungal promoters, along with a marker gene required for fungal selection. More recently, Cas9-sgRNA (single guide and tracrRNA) ribonucleoprotein complexes (RNPs) have been employed in a wide range of fungi [5
In an example of the RNP approach, targeted mutations have been introduced into the genome of the fungus Pyricularia oryzae
], which is among the most important fungal pathogens globally impacting cereal grain production. The fungus was transformed simultaneously with the RNPs to mutate the target gene and others to generate mutant genes that are positively selectable. This strategy is based on the presumption that those nuclei in which the genes are converted to their selectable forms are also those most likely to have taken up the RNP and consequently mutate the target gene as well.
We use Epichloë
spp. as exemplars of particularly difficult but important fungi for targeted genetic modification. Epichloë coenophiala
is widespread as a seed-borne endophyte of the highly popular pasture, forage, and turf grass, Lolium arundinaceum
(= Schedonorus arundinaceus = Festuca arundinacea
; tall fescue), although its existence was unsuspected in the first decades of widespread propagation of the grass during the mid-20th century. The fungus provides important benefits that translate to enhanced stand longevity and productivity and improved tolerance of drought and other stresses [9
], and it is capable of producing up to four different classes of alkaloids that protect the grass hosts against invertebrates [2
]. Unfortunately, the strains of E. coenophiala
that have been unwittingly co-propagated with tall fescue, and which remain dominant in much of the cool-season pasturelands, produce ergovaline, which is an ergot alkaloid of the highly toxic ergopeptine type [12
]. Levels of ergovaline tend to be very low, but they are often sufficient to at least cause reproductive problems and reduce livestock health and productivity. For the same reason, cultivars of Lolium perenne
(perennial ryegrass) with Epichloë hybrida
] strain Lp1 were pulled from the market in 1992 after it was determined that they had toxic levels of ergovaline [16
]. A study including deletion of the dimethylallyltryptophan synthase gene (dmaW
) from E. hybrida
Lp1 and its subsequent complementation with the ortholog from Claviceps fusiformis
] has demonstrated that dmaW
is essential for ergot alkaloid biosynthesis [19
]. There is a potential to develop and deploy such genetically altered strains of Epichloë
species in forage cultivars because the fungus can be cultured, manipulated, and reintroduced to produce new, stable symbioses with forage and pasture cultivars of tall fescue. However, it is desirable and perhaps essential that such modifications should not involve integration of any foreign gene in the genome, which is a requirement that makes the CRISPR RNP approach especially attractive.
The endophyte E. coenophiala
is a particularly difficult system for genetic inquiry because of its slow growth [20
] and the fact that it is a triploid interspecific hybrid [21
]. Two of its three ancestors were ergovaline producers, so E. coenophiala
has two homeologous copies of the ergot alkaloid biosynthesis (EAS
) gene clusters [2
]. An effort to delete the key gene dmaW
2 by marker exchange mutagenesis with a hygromycin B-resistance gene (loxP-flanked hph
) was a particular tour de force
in which over 1500 transformants were screened and the frequency of homologous gene replacement was 0.2% [22
]. Once a ∆dmaW
2 mutant was obtained and reintroduced into host plants, there was essentially no effect on ergovaline production because of the presence of its homeologue dmaW
1. Furthermore, although the selectable marker was readily removed from the ∆dmaW
2 mutant by transformation with a plasmid for the transient expression of Cre recombinase [22
], the resulting marker-free mutants consistently had lost the ability to establish stable symbiosis with tall fescue (unpublished results).
Natural mutants of Epichloë
species with deleted or inactivated alkaloid biosynthesis genes consistently have inactivated the entire set of genes for downstream steps [23
]. Whether or not this relates to the host incompatibility of the aforementioned marker-free ∆dmaW
2 mutants, the nature of these natural variants suggests that there is selection against expression of enzymes that, due to loss of upstream genes, no longer have access to their normal substrates. Therefore, we consider the most prudent approach to generating ergot alkaloid-negative mutants to be the deletion of both EAS
clusters in their entirety.
After genome sequencing revealed the subterminal location of the EAS
1 cluster in E. coenophiala
isolate e19, we devised a new approach to replace that cluster with a telomere-repeat array [24
]. Since the homeologous cluster, EAS
2, has an inactivating mutation in a late-pathway gene, lpsB
2, the resulting “EAS
1-knockoff” mutant produced only two early products of the ergot alkaloid pathway, chanoclavine and ergotryptamine. The technique employed transient expression of antibiotic resistance conferred by an hph
gene positioned in the vector to be lost subsequently by breakage of the integrated DNA at the introduced telomere repeat array. The success of this approach suggested that transient antibiotic selection could be used in other approaches for mutation. In this study, this strategy is applied to CRISPR-based deletion of both EAS
clusters as well as individual genes.
Both for research and for the practical aim of completely eliminating production of all ergot alkaloids from an agriculturally important grass symbiont, we have chosen to adapt a Cas9-sgRNA RNP approach to entirely eliminate both the 196-kb EAS1 cluster and the 75-kb EAS2 cluster. Here, we describe the success of that effort and follow-up experiments to demonstrate the facile nature of our approach and its broader applicability, opening the door to a wide range of non-transgenic manipulations of even slow-growing, asexual, polyploid fungi.
We have demonstrated that a CRISPR/Cas9 technology can be applied to polyploid Epichloë species for the precise removal of individual genes or gene clusters, and even the simultaneous removal of a pair of large clusters (196 kb and 75 kb). Specifically, the method utilized RNPs—consisting of sgRNAs and Cas9 protein that was translationally fused with two NLRs—which were introduced into the fungus by cotransformation with a transiently selected antibiotic resistance gene. There are two obvious advantages to this approach. One is that more than one gene or genome region can be deleted in a single procedure, and the other is that the procedure leaves no selectable marker or transgenes in the genome. The absence of the selectable marker in the final product allows for its reuse to eliminate additional genes or to reintroduce genes for complementation analysis, and it also addresses regulatory and public concerns about the use of transgenic organisms in applied research and agriculture.
We previously constructed a plasmid (pKAES329) to transiently integrate a selectable antibiotic-resistance gene (hph
) at a chromosome end [24
], and here, we have combined that approach with the RNP approach and found it to be successful and facile. However, we also questioned whether a transient integration of the marker was even necessary. We show here that marker integration is unnecessary by demonstrating the simultaneous elimination of two long gene clusters (EAS
1 and EAS
2) by treatment with the appropriate RNP mixtures together with a plasmid that, without integrating into the genome, provided expression of the selectable marker.
In this study, we did not test whether even transient marker selection was needed. However, we previously used the transformation of Epichloë
species without selection to delete a loxP-flanked hph
gene by transient expression of the Cre-recombinase gene [22
]. Although successful, the screen was tedious and time consuming, resulting in a 0.5–2.1% frequency of Cre-mediated deletions among the unselected colonies. In contrast, here, we report that plasmids mixed with the RNPs provided for temporary selection of a limited number of initially antibiotic-resistant transformants. From those, mutants with the target deletions were readily identified by PCR screens and a high proportion were marker free (18–100% depending on the experiment). Therefore, although integration of the plasmid-borne selectable marker was not required, our results suggest that that its inclusion in the transformation mixture aided in selection of the RNP-transformed isolates, including those with the desired deletions.
In targeting the EAS
1 gene cluster, a mismatch 3 bp upstream of the PAM site between the EAS2lpsBguide and its target genomic sequence near lpsB
1 did not prevent cleavage at this position. The mismatch at a single-nucleotide polymorphism between the sites near EAS
1 and EAS
2 (the sgRNA sequence was based on the latter) was in the “seed” region but not part of the “core” region for sgRNA-directed Cas9 cleavage [27
]. Since deletion of the 196-kb EAS
1 region was achieved with cleavage at the mismatch position, a precise match to the target was evidently not required.
Two additional genome alterations were observed in one of the ∆EAS1 ∆EAS2 mutants. One was the introduction of a telomere at the cut site near EAS1 in one of the mutants. Interestingly, that cut was repaired very simply with the addition of a single base pair followed by a telomere array. The other alteration in the same mutant was a recombination between the remnants of lpsA1 and lpsA2, presumably moving the genes near lpsA2 from genomic locations far from a telomere to close proximity to the newly generated telomere. Whether that alteration affects the expression of those genes may be an interesting topic for future inquiry.
Our results demonstrate the high efficiency of the CRISPR/Cas9 technology while not indicating any size limit for genome segments that can be efficiently and precisely deleted. The frequency of dmaW
2 deletion was 17%, compared to only 0.2% previously obtained for marker exchange mutagenesis of the same gene in the same E. coenophiala
]. In addition, the frequency of the 75-kb EAS
2 deletion (5.2%) was not dramatically less than that of the 2.6-kb dmaW
2 deletion (17%) and was very close to that of the 1.9-kb lolC
deletion (5.9%). Conceivably, size limits might be imposed by boundaries of chromatin states, so it is worth considering whether or not the clean deletions of large genome regions are more likely when those genes are part of a coordinately regulated cluster.
Though often characterized as “error-prone”, non-homologous end joining (NHEJ) frequently occurs without error. A study in mouse cell lines reported 70% precise NHEJ events after Cas9-mediated cleavage [28
]. Many target sites also exhibited approximately 10% 1-bp and 2-bp templated insertions. Insertions of 1 bp following Cas9 cleavage and NHEJ have also been described in Saccharomyces cerevisiae
, with 74–100% apparently being templated, implying that in those cases, Cas9 cleavage left a staggered (5′-overhanging) end that was filled in by DNA polymerase 4 [29
]. In our study, all of the eight sequenced junctions appeared to be precisely joined, but four of those had one or the other end cleaved 4 bp from the PAM site. In those four cases, the result was similar to the common 1-bp templated insertions observed by Lemos et al. [29
], so it is reasonable to speculate that they also resulted from staggered cleavage and end-repair.
As commercial sources have recently made Cas9-NLS fusion proteins and synthetic sgRNAs available, a variety of approaches that include Cas9-sgRNA RNPs have been employed in fungi. Most involve the integration of a stable selectable marker [30
] or simultaneous generation of a selectable mutation [8
]. Khan et al. [31
] report using the Cas9-sgRNA RNP system without selection to target the TOX3
effector gene in Parastagonospora nodorum
, with all of the six analyzed “transformants” exhibiting mutations at the repair site. This result contrasts with ours in that we found no mutations other than the deletions resulting from rejoining of the cleaved ends and, in approximately half of the junctions, an apparent 1-bp templated insertion. Thus, applications of this technology in different fungal systems may give substantially different outcomes.