Use of CRISPR cas9 for the genome editing of Fusarium graminearum, one of the main causal agents of Fusarium head blight

DIPARTIMENTO DI SCIENZE AGRARIE, ALIMENTARI E AGRO-AMBIENTALI

corso di laurea in BIOTECNOLOGIE VEGETALI E MICROBICHE

Use of CRISPR-Cas9 for the genome editing of Fusarium graminearum, one of the main causal agents of Fusarium Head Blight

RELATORE

Prof. Giovanni Vannacci Dr.ssa Sabrina Sarrocco

CORRELATORE

Prof. Tommaso Giordani

CANDIDATO Luca Malfatti

Anno Accademico

Per Aspera ad Astra

Index

1.3.1 Cas9-mediated gene knockout in filamentous fungi...17

1.4 Fusarium graminearum...19

1.4.1 Fusarium Head Blight...21

1.4.2 Potentiality of genome editing for the control of FHB...27

2. Aim of the work...31

3. Material and methods...32

3.1 Fungal isolate...32

3.2 Prediction of the spacer sequences (sgRNA) for targeted genome editing...32

3.3 Amplification of the RGR-cassette...33

3.4 Construction of the plasmid for targeted disruption of PKS12 in F. graminearum ITEM 124...36

3.5 Protoplasts preparation...40

3.6 Fungal transformation and transformants stabilization...42

4. Results and Discussion...44

4.1 Construction of the sgRNA-Cas9-expression plasmid...44

4.2 F. graminearum ITEM 124 Transformantion...52

5. Conclusions...61

Abstract

Fusarium graminearum Schwabe (teleomorph: Gibberella zeae (Schwein.)

Petch) is a plant pathogen that causes several diseases on small-grain cereals, such as seedling blight and foot rot, Fusarium Head Blight (FHB) and ear rot. FHB is one of the most economically worldwide devastating diseases of wheat (Triticum aestivum L.) and other cereal crops. In recent years the disease has caused extensive agricultural damage through direct losses in yield and quality due to the presence of Fusarium damaged kernels and their associated mycotoxins such as the trichothecene deoxynivalenol (DON). Despite of the availability of different strategies, such as crop rotation, tillage, the application of fungicides and the use of resistant cultivars or biopesticides, no one of these alone are able to control the disease.

The CRISPR-Cas (Clustered Regularly Interspaced Short Palindromic Repeats-CRISPR associated) technique has emerged in recent years as a powerful and fast-growing genome-editing technique and it is applied to different organisms thanks to its high efficiency, easy operations, and the possibility of multi-gene editing. The gene knock-out (KO) induced by the CRISPR-Cas9 is produced by small insertions or deletions (InDels) of nucleotides at the Double Strand Break (DSB) site which derives from the Cas9 endonuclease activity. In many published papers this technique was

applied to plants, animals and viruses, as well as in human medicine. Genome editing of filamentous fungi through the CRISPR-Cas9 technology has increased in recent years but, actually, very few reports about CRISPR-engineered filamentous fungi and only a minor number refers to plant pathogenic fungi.

The main purpose of this master thesis was to set up a protocol for the genome editing of a Fusarium graminearum mycotoxigenic isolate in order to demonstrate the applicability of the CRISPR-Cas9 technique on this organism. During the master thesis the genome of this isolate (ITEM124) was sequenced, annotated and released thus making easier all the bioinformatics work needed. A gene encoding a polyketide-synthase (PKS12) - which disruption is detected easily at a phenotypic level as slow growth and reduced sporulation - was chosen as target gene and used to design the RNA-guides to be included in the RGR-cassette. The cassette was then assembled in a Cas9 expressing plasmid containing also a marker gene (Hygromycin resistance) and a shortened AMA1 sequence, which allows to quickly removing the plasmid from the edited strain simply by reducing the selection pressure. The resulting vectors, one for each of the designed RNA-guide, were used for fungal transformation by protoplasts, these latter obtained according to a protocol optimized in the present work for our isolate. Putative mutants, resulting from different cloning and transformation experiments, were analyzed phenotypically and molecularly

in order to verify the knockout of the selected gene.

The ability to manipulate, at a genetic level, beneficial and plant pathogenic isolates with these technique represents a valid and promising tool to study how plant pathogens interact with their hosts as well as to improve the performance of beneficial isolates for the management of plant diseases.

Edited strains need to be checked for the presence of foreign DNA, to contribute to the debate about the exclusion of this type of genetically manipulated microorganisms from GMO.

1. Introduction

1.1 Genome Editing

Genome editing is a type of genome engineering in which DNA can be inserted, replaced, or removed from a genome using artificially engineered nucleases (Liu et al., 2015) and represents an useful tool to find out the function and effect of a gene or protein in a sequence-specific way (Burgess, 2013).

Genetically engineered and genome edited filamentous fungi are used as cell factories for a wide range of products of interest for various market sectors, from simple organic acids to complex secondary metabolites. In some cases – as for example for clinically significant drugs – their production was biotechnologically improved. The β-lactam group of antibiotics, including penicillin and cephalosporin, was the first to profit the progress made in molecular techniques for filamentous fungi. Other bioactive compounds produced by filamentous fungi, and important for the human health are cyclosporine A (immunosuppressive agent), lovastatin (cholesterol-lowering agent), taxol (anticancer agent) and griseofulvin (antifungal agent) (Meyer, 2008).

Genome editing protocols are applied also to fungi of interest in agriculture, both plant pathogens and beneficial isolates. Engineering fungal genome is a valid and promising tool to study how plant pathogens interact with their hosts as well as to improve the performance of beneficial

isolates for the management of plant diseases.

In the last years, Zinc-Finger Nucleases (ZFN) (Carrol, 2011) and Transcription Activator-Like Effector Nucleases (TALEN) (Cermak et al., 2011), are set up to facilitate precise genome modification. However, the time- and labor-intensive processes of using ZFN and the requirements of specific enzyme engineering for different targets in TALEN limit their applications. In this contest, the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/CRISPR-associated gene (Cas), in particularly CRISPR-Cas9 system II, is actually becoming the most popular genome-editing tool (Cong et al., 2013).

1.2 CRISPR-Cas

CRISPR (Clustered Regulatory Interspaced Short Palindromic Repeats) was first described in 1987 by Ishino and colleagues, when an “unusual structure” of the 3’ region flanking Escherichia coli iap gene was described to contain direct repeats of homologous sequences 29 nt in lengths, interrupted by 32 nt spacers. Anyway, the biological significance of the sequences was not known yet. This was followed by disparate descriptions of similar components in bacterial genomes, while any functional role of the encoded transcripts remained completely uncertain. Recognition that the CRISPRs harbour spacers that stem from virus or plasmids (Bolotin et al., 2005, Mojica et al., 2005, Pourcel et al., 2005), accompanied by the identification of associated cas genes products containing helicase and nuclease domains (Makarova et al., 2011) lead to the hypothesis that this system serves as mnemonic defense against foreign phages. In 2007, Barrangou and colleagues confirmed this idea in a study on Streptococcus

thermophilus.

CRISPR and CRISPR-Cas (Crispr associated proteins) constituting the CRISPR-Cas system have been divided into three types (I, II and III) (Haft et al., 2005, Makarova et al., 2011). Compared with types I and III, which possess more complex molecular mechanisms and need multiple Cas proteins working together to cleave the target DNA, the type II CRISPR system (Figure 1.1) from Streptococcus pyogenes resulted much simpler

and was consequently been applied widely (Makarova et al., 2013, Chylinski et al. 2013). This system consists of only two elements: a Cas9 nuclease and a single-guide RNA (sgRNA), which is composed of two small RNAs, a target-recognizing CRISPR-RNA (crRNA) and auxiliary noncoding trans-activating crRNA (tracrRNA) (Jinek et al., 2012, Kuscu et al., 2014). The idea at the base of the type II system is that the RNAs encoded by CRISPR locus stem from viral encounters form a complex with the cas-encoded protein to guide an endonuclease to homologous sequences within the host, where DSB are induced to interfere with viral proliferation (Makarova et al., 2011).

Figure 1.1: Type II CRISPR system.

The principle is that CRISPR-Cas-mediated immunity of bacterial hosts against invading viruses is sequentially executed by three distinct processes, which are generally referred to as acquisition, expression, and interference phase (Bondy-Denomy et al., 2014). In the acquisition phase DNA fragments from previously encountered phages or plasmids are integrated and stored as spacer regions in the CRISPR array to expand the host memory of encountered invading pathogens. In case of a further

infection the corresponding spacer region is ready to become part of a processed crRNA that, with a tracRNA, is incorporated in a complex with the Cas proteins. Thanks to the complementary sequence homology of the crRNA, this complex is then directed to the target site where cleavage is executed. Recognition and processing depend on the complementarity of the spacer with its target site, which cover the protospacer sequence and an adjacent motif (PAM: Protospacer Adjacent Motif) at its 3’ end. This motif prevents the retargeting of the system to the CRISPR array but it is essential for driving the Cas9 activity to its non-self template (Jinek et al, 2012, Doudna et al., 2014, Bondy-Denomy et al., 2014, Chylinski et al 2014, Gasiunas et al., 2012). In recent years, CRISPR-Cas system has emerged as a potential candidate to solve the problem of low gene editing frequency in filamentous fungi (Tian-Qiong et al., 2017). CRISPR-Cas9 has become one of the fastest-growing gene-editing technologies and is applied to various species including filamentous fungi thanks to significant advantages such as high efficiency, easy operations, and the possibility of multi-gene editing, just to mention some. The CRISPR-Cas9 system therefore has become a further powerful genome-editing technology in addition to Zinc Finger Nucleases (ZFN) and Transcription Activator Like Effector Nucleases (TALEN) (Mashimo, 2014, Wu et al., 2015, Lee et al., 2016, Estrela et al., 2016).

1.3 CRISPR-Cas in fungi

Since the first DNA transformation of a filamentous fungus (Neurospora

crassa) in 1973 by Mishra and Tantum, genetic engineering technologies

opened the door for basic biological research in filamentous fungi. The development of molecular-biology research in these organisms has grown rapidly thanks to a new variety of technical methods such as encompassing RNA interference, gene targeting, in vitro transposon tagging, heterologous expression, and gene knockout. These methods have allowed scientists to investigate and exploit biosynthetic and regulatory mechanisms in filamentous fungi (Weld et al., 2006, Kück et al., 2010, Jiang et al., 2013).

Furthermore, with the advent of high-throughput sequencing technologies, the number of sequenced genomes of filamentous fungi has also been rapidly increasing, and they further revealed the existence of a large number of uncharacterized and silent secondary metabolite gene clusters (Andersen et al., 2013).

Figure 1.2: Homologous Recombination (HR) and Non-Homologous End Joint (NHEJ) mechanisms

Genome engineering techniques that use Site-Specific Nuclease (SSN) like Crispr-Cas9, induce a targetable DNA double-strand break (DSBs). DSBs of DNA are also common responses due to the natural mechanical stress such as exposure to ionizing radiations, radiometric chemicals, transgenic DNA, or crossing over during meiosis division (Britt, 1999). Homologous Recombination (HR) and Non-Homologous End Joint (NHEJ) (Figure 1.2) are the two common endogenous mechanisms for the repairing of DNA DSBs. HR is a precise, trustable, but slow, repairing mechanism that uses homology templates or an exogenous DNA fragment for target gene

modification or gene insertion. HR repair of DSBs mostly occurs at S or G2 stages of the cell cycle during the contiguity of sister chromatids (Bortesi et al., 2015).

Unlike HR, NHEJ is active throughout the cell cycle and has a higher capacity for repair, as there is no need for a repair template (like sister chromatid, homologue, or exogenously provided DNA), it is the major frequent pathway (Sonoda et al., 2006), and it is faster than HR. However, NHEJ is an error prone repair system that depends on random insertions or deletions (InDels) of nucleotides, causing frame shift mutations thus effectively creating gene knockout (Kim and Kim., 2014, Steentoft et al., 2011). These InDels generated during repair by NHEJ are typically small (1-10 bp) but extremely heterogeneous. NHEJ is the most common DSB repairing mechanism in microorganisms, comparing to the lower target integration by HR (Puchta, 2005). Three enzymatic activities are essential for NHEJ: nuclease to cut the damaged nucleotide, polymerase to repair, and ligase to restore the phosphodiester bond (Lieber, 2008). In conclusion, NHEJ is the most conserved mechanism in fungi for chromosomal DSBs (Carvalho et al., 2010).

From a molecular point of view, HR repair depends on Rad52 proteins that mediate targeted integration between two homologous DNA sequences, while the NHEJ pathway is mediated by activities of Ku heterodimer (Ku70/Ku80-protein complex) and DNA ligase IV to ligate DSBs without

any homology (Shrivastav et al., 2008). Inactivation of Ku70/Ku80 proteins as a component of NHEJ mechanism in Neurospora crassa induces the frequency of the HR mechanism up to 100% (Meyer, 2008). Similarly, knockout of akuA gene in A. fumigatus encoding Ku70 protein that mediates the NHEJ indirectly promotes the HR repairs (Krappmann et al., 2006).

1.3.1 Cas9-mediated gene knockout in filamentous fungi

At the beginning, for the CRISPR-Cas9 technique on filamentous fungi, RNA polymerase III promoters were used for the transcription of sgRNA. In 2015 Nødvig and colleagues adopted a new strategy for the release of the sgRNA module. The authors fused the HH (Hammer Head) and HDV (Hepatitis Delta Virus) ribozyme on the 5’ and 3’ ends of the sgRNA sequence. Through self-processed RNA cleavage and subsequent release of the gRNA sequence without modifications, CRISPR-Cas9 was successfully implemented in six different fungal species. This method has been demonstrated to be valid in filamentous fungi and was used in Phytophtora

sojae, Aspergillus fumigatus, Talaromyces atroroseus, and A. carbonarius

(Fang and Xia., 2015, Weber et al., 2016, Nielsen et al., 2017, Weyda et al., 2017). Albeit the gene editing efficiency varied extremely between different fungi (from 1 to 100%), the advent of this approach appears to have solved the sgRNA constructions problems, which should greatly accelerate the development of CRISPR-Cas9 in filamentous fungi (Tian-Qiong et al., 2017).

Fuller and colleagues (2015) described the NHEJ-mediated integration of transforming DNA into Cas9 cleavage site. Commonly, filamentous fungi have a dominant NHEJ repair pathway, which results in random insertions or deletions of one or more nucleotides that can lead to gene disruption after formation of a DSB (Zhang et al., 2015). Nonetheless, in the A.

fumigatus Cas9 knockout system Bachu and colleagues (2015) found that

DNA from phenotypically positive colonies shows a premature translational stop and the larger insertions of the transforming DNA at the Cas9 cleavage site. In a recent study Gardiner and colleagues (2017) found that the MMEJ (Microhomology-Mediated End Joining) DNA repair pathway is likely to be the predominant process involved in erroneous repairing of Cas9-induced DSB in F. graminearum. The authors also demonstrated that is possible to edit the genome of F. graminearum via CRISPR-Cas9, with a homologous recombination of a vector containing everything necessary to editing the genome in the genome itself, without the need for any expression plasmid. In this case the exogenous DNA remains in the genome of the fungus.

In addition, a new article was recently reported in Science to introduce the application of the CRISPR-Cas9 system in the higher fungus Agaricus

bisporus. By deleting one of the six polyphenol oxidase (PPO) genes in the

mushroom’s genome, the activity of PPO decreased by 30%, which effectively mitigated the browning of A. bisporus. Although this news does not disclose the experimental process, the CRISPR-Cas9 system was proved to be effective in higher fungi for the first time (Waltz, 2016, Yin et al., 2017).

1.4 Fusarium graminearum

Fusarium graminearum Schwabe (teleomorph: Gibberella zeae (Schwein.)

Petch) is a common plant pathogen that causes several diseases on small-grain cereals, such as seedling blight and foot rot, Fusarium Head Blight (FHB) and ear rot (Popovsky et al., 2012).

Fusarium graminearum species includes isolates mostly belonging to

Group 2, whose members are discerned from the morphologically similar heterothallic form, named Group 1. While the isolates of both Groups could produce FHB symptoms on wheat, only Group 1 isolates produce typical symptoms of crown rot and foot rot of wheat, oats and Medicago spp. (Purss, 1971). From 1999 both Groups can be distinguished on the basis of molecular, morphological and cultural characters (Aoki et al., 1999). Group 1 was recognized as F. pseudograminearum, and its teleomorph was described as Gibberella coronicola. The name F. graminearum and its associated G. zeae teleomorph were retained from Group 2. Analysis of six genes among F. graminearum isolates revealed seven phylogenetically distinct lineages (O’Donnel et al., 2008) that where biogeographically structured. The identification of congruent and apparently discrete groups permits successive study to determine if these groups differ significantly in biological characteristics that impact on their ecology and disease control (Xu et al., 2009).

with a velvety hairy aerial mycelium (Figure 1.3). Colonies pigmentation varies from pale rose and brown to bluish violet depending on species and growth conditions. In particular F. graminearum colonies pigmentation varies from light reddish purple to light orange, while on corn, it varies from white to pink (Batt et al., 2014).

Figure 1.3: Fusarium graminearum colony on PDA.

The typical Fusarium spore (macroconidium) is fusiform, multicelled by transverse septa and has the characteristic foot-shaped basal cell together with a wisp-like apical cell (Goswami et al., 2004).

although soil, seeds and different hosts are also inoculum sources (Sutton, 1982). Its colonies overwinter on crop residues of maize, wheat and barley and form both perithecia, that will produce ascospores (sexual spores) and macroconidia (asexual spores). Ascospores are blown or splashed to the new site of infection (wheat flowers) at the beginning of the favourable season and develop the primary infection (Leplat et al., 2013). F.

graminearum has an optimum temperature for growing at 28-29°C (Xu et

al., 2009).

1.4.1 Fusarium Head Blight

Fusarium Head Blight (FHB) or scab, is one of the most economically worldwide devastating diseases of wheat (Triticum aestivum L.) and other cereal crops that causes extensive yield and quality losses in humid and semi-humid regions (Guenther, 2005, Palazzini, 2015). This disease (Figure 1.4) was first described in 1884 in England and was considered a major threat to wheat and barley during early years of the 20th century (Muriuki, 2001, Stack, 1999, 2003) FHB has been identified by CIMMYT (Centro Internacional de Mejoramiento de Maiz y Trigo) as a major factor limiting wheat production in many parts of the world (Stack, 1999), such as North and South America, Canada, Europe, and Asia (Goswami et al., 2004, Chakraborty et al., 2006, Tunali et al., 2012).

Figure 1.4: Fusarium Head Blight (FHB) disease cycle.

FHB has been associated up to 19 species, but the most prevalent pathogens are F. graminearum, F. culmorum, F. avenaceum, F. poae,

Microdochium nivale and M. majus (Dill-Macky et al., 2000, Xu et

Nicholson, 2009, Popovski et al., 2013). In Italy, FHB has been detected for several years, indeed many environmental monitoring in the main areas of cereals production have been confirming the incidence of the main causal agents of the FHB (Figure 1.5). The incidence is closely related to climatic conditions and cultivation areas. The presence of F. graminearum is higher in Northern and Central Italy, while in the South the presence of

F. culmorum is higher (Pasquini et al., 2006).

The composition, the development, and the structure of the Fusarium community depend on a combination of factors, among which climatic conditions play a major role (Muller et al., 2010). For instance, Ramirez and colleagues (2006), found that the mycelial growth of two strains of F.

graminearum reached an optimum at 25°C at water activities ranging

between 0.950 and 0.995, and that no growth was observed below 5°C. Both strains were able to grow in dry conditions at a minimum water activity of 0.900.

Wheat residues, like straw, are the main source of the primary inoculum for the causal agents of FHB. The main part of the FHB species life cycle is saprotrophic and counts on maintaining the occupation of colonized plant debris in competition with other microorganisms (Harris et al., 1999).

Figure 1.5: Symptoms (premature yellowing of spikes) of Fusarium head blight on wheat.

FHB not only causes quantitative yield loss but may also produce quality problems of the grains because of the production of mycotoxins. Mycotoxins are capable of inducing both chronic and acute toxic effects in animals depending on the type, the level and the duration of exposure, the animal species that is exposed, and the age of the animals (Gunther et al., 2014). In FHB most mycotoxins belong to the class of trichothecenes. Deoxynivalenol (DON) and its acetylated derivatives (3- and 15-acetyl-DON) are the most produced molecules of this class (Harris et al., 1999). All these molecules are Type B trichothecenes (Starkey et al., 2007), sesquiterpenes with a low molecular weight and volatile (Rocha et al., 2005).

Common effects of trichothecene toxicity in humans are depression of immune responses and nausea, sometimes vomiting (DON is also called Vomitoxins for this reason) (Peraica et al., 1999, Windels, 1999).

Different strategies are used to reduce the impact of FHB like crop rotation, fungicide application, tillage practices and of course the use of less susceptible cultivars. Among all these strategies, fungicide control seems to be the most efficient (Homodork et al., 2000, Mastherázy et al., 2011), even if it was observed that some types of fungicides could boost DON content on grains (Ramirez et al., 2004) and pathogens can generate fungicide resistance (Yuan, 2005).

In this context, biological control by the use of beneficial organisms appears to be very promising. Because this pathogen is generally considered a poor competitor over time compared to other organisms that colonize crop residues (Pereyra and Dill-Macky, 2008), competition for cultural debris could thus be a valid strategy to control the production of the primary inoculum of the causal agent of FHB (Leplat et al., 2013). If biocontrol agents are inoculated on cultural debris in soil, they can access to territory or resources previously held by the pathogens (Jensen et al., 2016). As they are able to outcompete with this pathogen, many fungi are studied in terms of their ability to limit the survival of F. graminearum. Of these, Trichoderma atroviride, Trichoderma harzianum and Clonostachys

to colonize wheat residues or to reduce F. graminearum sporulation on cultural debris (Gromadzka et al., 2009; Sarrocco and Vannacci, 2017).

Trichoderma gamsii T6085 has been used by the Plant Pathology and

Mycology Lab of the Department of Agriculture, Food and Environment (University of Pisa) both in laboratory and field experiments for the control of FHB causal agents, and has reduced F. graminearum and F. culmorum growth by acting as an antagonist and a mycoparasite, also reducing deoxynivalenol content (Matarese et al., 2012; Sarrocco et al., 2013; Baroncelli et al., 2016).

Among biocontrol agents of FHB, bacteria such as Bacillus and

Streptomyces (Palazzini et al., 2018), Pseudomonas (Khan et al., 2009) and Lysobacter (Jochum et al., 2006) have been used. Also, Cryptococcus

species have shown antagonistic activity towards FHB pathogens. Biocontrol of pathogens can be achieved through antibiosis, mycoparasitism, competition, and the induction of resistance in the host plant (Legrand et al., 2017).

1.4.2 Potentiality of genome editing for the control of FHB

The use of the CRISPR-Cas9 system for the genome editing and the capability to insert, delete and replace target genes can generate positive multiple gene modifications in fungal pathogens (Cong et al., 2013) like

Fusarium graminearum. The target of these mutations could be the genes

for polyketide synthase (PKS), an essential enzyme for the biosynthesis of toxic compounds. Therefore, these modified filamentous fungi cannot produce toxic compounds, which significantly reduce the fungi harmfulness to the host (Fuller et al., 2015). For instance Malz and colleagues (2005) found that the mutant of F. graminearum for the PKS12 gene showed no transcript of the PKS12 gene and was unable to produce aurofusarin, a dimeric polyketide belonging to the aromatic group of polyketides, the so called naphtoquinones, which are toxic compounds especially for poultry.

Strains of F. graminearum unable to produce mycotoxins could be used as biocontrol agents against mycotoxigenic Fusarium wild type strains. For instance, in the biological control of aflatoxin contamination of crops the greatest successes have been achieved using non-toxigenic strains of

Aspergillus flavus and A. parasiticus. Aspergillus species are generally

found in soil and crop debris, which acts as the principal source of primary inoculum for infecting maize (Jaime-Garcia and Cotty, 2004). Atoxigenic (or nontoxigenic) isolates, defined as isolates that do not produce any

aflatoxin, of A. flavus have been used as biological control agents to compete with and displacing aflatoxin producers (Atehnkeng et al., 2008) Applications in both pre- and post-harvest of atoxigenic strains of

Aspergillus species, especially with peanuts and cotton, have shown

considerable reductions in aflatoxins contaminations between 70% to 90% (Dorner, 2004, Pitt and Hocking, 2006, Dorner, 2008). Zanon and colleagues (2015) found that inoculum of peanuts in soil field with two atoxigenic strains of A. flavus reduced dramatically the aflatoxin contamination around 86%.

Atoxigenic strains competitively block naturally toxigenic strains in the same niche and contend for crop substrates. For this reason atoxigenic strains must be predominant in the agricultural environment when the crops are susceptible to be infected by the toxigenic strains. Field tests showed that atoxigenic strains reduced naturally Aspergillus populations up to 99% in the soil of peanuts fields (Yin et al., 2008).

While the US authorities have previously expressed that crop varieties made through genome editing do not constitute Genetically Modified Organisms (GMOs) (Jones, 2015), in the European Union the legitimacy of the use of organisms edited by Crispr-Cas9 is unclear: an organism edited with Crispr-Cas9 in which it has been added an exogenous DNA falls into the GMOs legislation, while an organism in which the only modification is a deletion of a sequence of DNA falls in a legislative

lacunae.

The European GMO Directive 18/2001/EC regulates the deliberate release into the environment of GMOs and their placing on the market within the EU, the organisms covered by that Directive must be authorised after an environmental risk assessment. The Directive does not however apply to organisms obtained through specific techniques of genetic modification (Bobek, 2018). The Directive defines GMO in Article 2(2): “An organism, with the exception of human beings, in which the genetic material has been altered in a way that does not occur naturally by mating and/or natural recombination”. According to this definition a GMO must be “genetically modified”, which means that it must be an inheritable modification of genetic material that could not have appeared naturally by mating or natural recombination. Such modification of genetic material consists in the addition of foreign genes into the organism (Glas and Carmeliet, 2017). In 2015 the European Academies’ Science Advisory Council (EASAC) asserted that the products of New Breeding Techniques (like Crispr-Cas9) should not occur under GMO legislation when they do not have foreign DNA (EASAC 2015).

Advocate General Michal Bobek in his opinion of 18 January 2018 considers that an organism obtained by mutagenesis can be a GMO if it satisfies the substantial criteria enforced in the GMO Directive and he observes that that Directive does not require the addition of foreign DNA in

an organism in order for the latter to be characterised as a GMO, but simply says that the genetic material has been modified in such a way that does not come out naturally. In conclusion, he says that mutagenesis techniques are free from the obligations of the GMO Directive providing that they do not include the use of recombinant nucleic acid molecules or GMOs other than those produced by one or more of the methods listed in Annex I B (Bobek, 2018)

2.

Aim of the work

Aim of the present work was to set up a genome editing protocol, by the use of CRISPR-Cas9 approach, as a proof of concept of the applicability of this technique on Fusarium graminearum. Target of the silencing was the

PKS12 gene, a gene encoding for a polyketide synthase whose deletion can

cause modification at a phenotypic level. In order to set up this genome editing protocol, different RNA guide sequences have been identified and used to create a RGR cassette to be assembled in final plasmids, also containing the CAS9 encoding gene and a marker gene (Hygromycin resistance), to be used for fungal transformation.

In order to obtain silenced mutants, a protoplast preparation protocol was defined as well as a transformation method.

3. Material and methods

3.1 Fungal isolate

This study was conducted on the mycotoxigenic strain Fusarium

graminearum ITEM 124, also known as ATCC 56091, isolated in 1976

from rice harvested in Vercelli, Piemonte, Italy. F. graminearum ITEM 124 was kindly provided by ISPA, CNR, Bari.

This isolate was conserved on PDA (Potato Dextrose Agar, Difco, CA, USA) under mineral oil at 4°C and actively grown on PDA at 25°C with photoperiod consisting of 12h light/12h darkness in a thermostatic cell. The genome of the isolate was recently sequenced, annotated and released, as described in Zapparata et al. (2017).

3.2 Prediction of the spacer sequences (sgRNA) for targeted genome editing

The target gene PKS12 was used to predict the spacer sequences by using Benchling, a free web based tool (https://benchling.com/). Benchling allows to assess for on-target efficiency by algorithm built into the online program (Doench et al., 2016), to which 20 bp long spacers are calculated to have a score >70. These 20 bp long spacers, here named spacer N1-20_,

were predicted to use Cas9 mediated DSB within coding region of the target gene. The N1-20 _{spacer has to be adjacent to a motif (PAM:}

Protospacer Adjacent Motif) at its 3’ end. This motif prevents the retargeting of the system to the CRISPR array but it is essential for driving the Cas9 activity to its non-self template.

3.3 Amplification of the RGR cassette

The Ribozyme-sgRNA-Ribozyme (RGR) cassette (Figure 3.1), containing the M6-1_{, the Hammer Head ribozyme, the spacer N}1-20_{, the chimeric}

tracrRNA and the HDV ribozyme (211 bp total), was constructed by inserting the spacer sequence into the sgRNA, containing a PacI restriction site flanked by gpdA promoter and TrpC terminator sequences originating from Aspergillus nidulans.

Figure 3.1: RGR cassette used to create the final deletion plasmid. The cassette contains the M6-1_{, the Hammer Head ribozyme (HH), the spacer N}1-20_{, the chimeric}

tracrRNA and the HDV ribozyme

For each spacer, the RGR-cassette was altered through two separate rounds of site-directed mutagenesis (SDM).

First round SDM PCR was performed using the plasmid pCRISPRa1.1 as template with primers F2 (one for each spacer) and Universal Reverse (5’-GACATGGAGCTATTAAATCAGTCCCATTCGCCATGCCGAA-3’), simultaneously modifying the 20 bp spacer sequence and inserting a

downstream adapter onto the amplicon. The sequences of F2 primers are shown in Results.

The PCR protocol was as follows:

Cycle Step Temperature (°C) Time Cycles

Inital Denaturation 98 30'' -Denaturation 98 8'' Annealing 70 30'' 24 Extension 72 13'' Final Extension 72 5' -Hold 4 ∞

-The PCR was performed by using a HiFidelity Q5 Taq Polymerase (Biolabs, England) in 50 L final volume, according to the manufacter’s instruction.

The pCRISPRa1.1 (Figure 3.2, showing the RGR-cassette used as template for building the RGR) contains the RGR-cassette, with the two ribozymes (HH and HVD) and the tracrRNA protospacer, without the spacer sequence

Figure 3.2 RGR-cassette included within the pCRISPRa1.1 plasmid and used as template for the construction of the RGR-cassettes.

The PCR products were purified on 1% agarose gel (QIAquick Gel Extraction Kit, Qiagen, Hilden, Germany, according to the manufacter’s instruction) and used as template for the second round of SDM PCR where primers F1 (whose sequences are shown in Results) and Universal Reverse were used to mutate the six first nucleotides (M6-1_{) at the beginning of the}

5’-end of the HH ribozyme and to incorporate the upstream adapter for the final assembly. The PCR protocol was the same as described above. The protocol used to create the RGR-cassette is shown in Figure 3.3.

Figure 3.3 Scheme of the PCR protocol that allows to insert the spacer into the RGR-cassette starting from the p1 (pCRISPRa1.1) plasmid and using primers F2 and F1 designed on the basis of the N1-20 _spacers.

3.4 Construction of the plasmid for targeted disruption of PKS12 in F. graminearum ITEM 124

The final plasmids (one for each sgRNA) were generated by inserting the respective sgRNA fragments into the pFC332 plasmid (Figure 3.4). The plasmid contains not only the gene coding for the Cas9 protein, but also the marker gene hph, conferring resistance to Hygromycin and the AMA1 gene, that allows the auto-replication of the plasmid when under pressing condition (antibiotic into the growth medium).

PCR 1

PCR 2

Figure 3.4: Map of the pFC322 plasmid containing the Cas9, the AMA1 and the Hygromycin (hph) resistance gene.

The Cas9 expressing plasmid was digested with PacI restriction enzyme for 2 hours at 37° and heat-inactivated at 65°C. The linearized pFC332 plasmid was assembled with each of the final RGR amplicons (211bp) in a 1:3 molar ratio in a 10 µl HiFi reaction (Gibson et al., 2010), by using the NEBuilder HiFi DNA Assembly Cloning Kit (New England BioLabs), according the manufacter’s instruction. The complete scheme for the deletion plasmid construction is in Figure 3.5.

Figure 3.5. The process of spacer sequence insertion is based on two forward primers

and one reverse primer. (The Forward primers are different for each of the spacers N 1-20_{: M}6-1 _{is reverse complementary to N}1-6_{). The expression vector is linearized by PacI} digestion. The final plasmid is assembled (Gibson et al., 2010)

The assembly reaction was incubated for 15 min at 50°C and afterwards sub-cloned directly into competent E. coli cells DH10β (New England Biolabs), according to manufacturer’s protocol.

The success rate of mutation and assembly was usually higher than 90% but as a precaution, the final constructs were checked by colony PCR and sequencing to confirm they were error free.

The colony PCR was performed by using universal primers SB011

(5’-ACTCCATCCTTCCCATCCCTTA-3’) and SB012

Cycle Step Temperature (°C) Time Cycles Inital Denaturation 96 30'' -Denaturation 95 45'' Annealing 52 60'' 25 Extension 72 30'' Final Extension 72 5' -Hold 4 ∞

-The PCR was performed by using a Taq DNA Polymerase with standard Taq Buffer (Biolabs, England and according to manufacter’s instruction) in 50 L final volume, according to the manufacter’s instructions using directly a small amount of the E. coli colony as template.

The amplicons from the new constructs should contain 211 bp more than the amplicon obtained by using pFC322 (326 bp) as template, and they were run in a 2% agarose gel and using a low Molecular Weight Ladder (Biolabs, England). The amplicons were sequenced at the BMR genomics (Padova, Italy) and the sequences compared with that from plasmid pFC332 and from the expected 211 bp sequence of the RGR cassette. Those plasmids containing the right construct were isolated from E. coli by using the Mini Prep Qiagen kit (Qiagen, Hilden, Germany), according to the manufacter’s instructions and used for the fungal transformation.

3.5 Protoplasts preparation

Different protocols were tested in order to obtain an enough number of fungal protoplasts of good quality and able to regenerate. The following protocol resulted to be the most adapt for our isolate.

Protoplasts of F. graminearum ITEM124 were prepared by inoculating a spore suspension (4·105 _{conidia mL}-1_{) of the isolate on a sterilized}

cellophane disc placed into a PDA plate and incubated at 25°C, 12h light/12h darkness cycle for 24 hour until the development of the germ tubes. Then the cellophane containing the isolate was transferred in an empty plate and covered with 5 mL of the Lysing solution. The plate was incubated with slight oscillation at 28°C.

The Lysing solution was made of:

 NovoZymTM (Novo Biolabs) [20 mg mL-1]: 50% ;  Cellulase (Sigma) [100 mg mL-1]: 10% ;

 Driselase (Sigma) [50 mg mL-1]: 20% ;  Buffer magnesium sulfate: 20%.

All enzymes were dissolved in the magnesium sulfate buffer, made by:  MgSO4 1.3M;

 NaH2PO4 1.0M;

 Na2HPO4 1.0M.

After 30 minutes the disc was washed with the Lysing solution and removed from the Petri dish, the digestion went on until the digestion of the

mycelium and the development of protoplasts (3-4 hours) under the same conditions previously described. Protoplasts formation was microscopically monitored after 2 hours of incubation, every hour until the release of protoplasts due to mycelium digestion.

The digestion solution, containing protoplasts, was filtered by glass wool in a Falcon tube thus holding undigested mycelium and letting protoplasts to flow out. 1 mL of magnesium sulfate buffer was used to wash the wool in order to collect the remaining protoplasts.

The filtrate was aliquoted in 1.5 mL Eppendorf tubes and protoplasts were pelleted by centrifuging for 10 minutes, 3000 rpm at 4°C. The supernatant was carefully discarded and the pellet re-suspended in 1 mL of STC. This step was repeated three times and in the latter the pellet was re-suspended in 600 μL of STC buffer + 150 μL of 40% PEG. 200 μL of the final suspension was transferred in cryogenic tubes and stored at -20°C until transformation. STC buffer (100 mL):  Sorbitol: 18.4 g;  TRIS buffer: 5 mL;  CaCl2: 5 mL;  H2O: till 100 mL.

TRIS buffer was prepared by dissolving 6.05 g of TRIS Base in 50 mL of H2O.

The viability and the regeneration ability of protoplasts were checked by plating an aliquot on PDA containing sorbitol 1M and incubating plates at 25°C, 12 hours light/darkness.

3.6 Fungal transformation and transformants stabilization

Transformation was carried out by adding 1 to 10 μg of purified plasmid DNA to 200 μL of protoplast suspension, which was subsequently incubated on ice for 15 minutes. Then, 1 mL of 40% PEG was added and incubated for 15 minutes at room temperature. The suspension was added to 12 mL of soft PDA (8 g L-1 _{agar) containing sorbitol 1.0 M and 200 ppm}

of Hygromycin B. The medium was poured on Petri dish containing a layer PDA 1M sorbitol (20 g L-1 _{agar). The plate was incubated at 25°C until the}

transformants appeared. As soon as Hygromycin resistant colonies developed into the plates, they were immediately transferred on new PDA plates containing 200 ppm of Hygromycin and incubated at the same conditions, then transferred for three following times on PDA without antibiotic (no selective pressure) in order to check the morphology of the colonies (if slow growing and without red pigments as expected). All putative transformants were then confirmed by diagnostic PCR and sequencing analysis. Primers for deletions analysis were designed to bind approximately 380bp upstream and downstream of the expected double-stranded breaks. For the deletion occurring at 1001 bp primers

PKS12_799F (5’-AGTGCCGATCTATGCTCCGTAC-3’) and PKS12_1184R (5’-ACTCGGTTTGTGTCTCTGCTGA-3’) were used, whereas for deletion at 1326 bp PKS12_1296F ACAAAGGTCGCGATGTTCATCG-3’) and PKS12_1685R (5’-TGCCACCTGTAGTCTGCAATCA-3’) were used. PCR protocol, performed in 50 L final reaction, was made by using Taq DNA Polymerase with standard Taq Buffer (Biolabs, England and according to manufacter’s instruction) was as follows:

Cycle Step Temperature (°C) Time Cycles

Inital Denaturation 944 5’ -Denaturation 94 1’ Annealing 65 1’ 34 Extension 72 3’ Final Extension 72 10’ -Hold 4 ∞

-PCR products were sequenced at BMR (Padova, Italy) and compared with the sequence form the Wild Type (WT) (obtained using primers binding approximately 380bp upstream and downstream of the sequences used a spacer N1-20_{) in order to detect the deletion due to Cas9 activity.}

4. Results and Discussion

4.1 Construction of the sgRNA-Cas9-expressoion plasmid

We managed to construct the RGR-cassette and inserted it in the Cas9 expression plasmid pFC332 (Nødvig et al., 2015). To get this we started by choosing two high-on target-efficiency spacer sequences within the domain through the analysis of the genome of our F. graminearum isolate (ITEM 124, accession number NQC00000000, BioProject PRJNA397367; BioSample SAMN07455307), recently sequenced, annotated and released (Zapparata et al., 2017).

The choice of the PKS12 gene was due to the fact that this gene is present in the aurofusarin cluster and it is responsible for the production of the aurofusarin pigment. The silencing of this gene leads to white-colony mutant (Frandsen et al., 2006) easy to be phenotypically distinguished from the WT.

The on-target efficiency regards the probability of the gRNA to target the gene sequence. Several spacer sequences were assessed for on-target efficiency (Doench et al., 2016) by algorithm built into the online program Benchling (http://www.benchling.com), to which 20 bp long spacers, here named spacer N1-20_{, can be calculated and chose among those having an}

After the bioinformatics analysis two spacers were identified, named 1001 and 1326 – within the PKS domain (Figure 4.1), that predicted to direct Cas9 cleavage to 1001 nucleotides downstream of the start codon (PKS12_1001 spacer sequence: 5’-GAGACCTTGGTAAGATCCAG-3’) and to 1326 downstream of the start codon (PKS12_1326 spacer sequence: 5’-TGTTCATCGACAAATTCCCG-3’).

Figure 4.1: 1001-N1-20 _{and 1326-N}1-20 _{spacer within the PKS domain.}

These spacer sequences were used to design the F2 and F1 primers for each of the two targets (Figure 4.2). These primers were designed on the bases of each gRNA sequence and consist of 60 bp. The primer F2 have the sequence of one fragment of the HH ribozyme and also the sequence of all the tracrRNA, which are invariable, so in this primer we added the spacer sequence between the HH and the tracrRNA. In the F1 we have the other fragment of the HH ribozyme and the PacI restriction sites in order to ligate the cassette to the Cas9 expressing vector linearized with PacI, so in this primer we added the reverse complement of the six first bases (M6-1_{) of}

the spacer between the HH and the PacI restriction site. In this way the target gene is defined to be cleaved by Cas9, thus conferring the specificity of the CRISPR-Cas9.

Figure 4.2: 60bp primers F1 and F2 designed on the basis of the N1-20 _{spacers detected}

within the PKS12 domain. At each color of the bases correspond a different part within the final RGR cassette.

With the Gibson cloning method we easily built the Ribozyme-sgRNA-Ribozyme (RGR) cassette inserting the spacer sequence into the sgRNA (included within the pCRISPRa1.1 plasmid, used as template for the first round of PCR) with only two PCRs.

The final schemes of the two RGR-cassettes are shown in Figure 4.3. The cassettes include:

- M6-1_{: reverse complement sequence of the Hammer Head ribozyme;}

- HH: sequence of the Hammer Head ribozyme; - N1-20_{: gRNA within the target gene;}

F1_RGR_PKS12_1001: 5’-CGCTTGAGCAGACATCACCGGGTCTCTGATGAGTCCGTGAGGACGAAA CGAGTAAGCTC -3’ F2_RGR_PKS12 _1001: 5’-ACGAAACGAGTAAGCTCGTCGAGACCTTGGTAAGATCCAGGTTTTAGAG CTAGAAATAGC-3’   F1_RGR_PKS12_1326: 5’-CGCTTGAGCAGACATCACCGTGAACACTGATGAGTCCGTGAGGACGAA ACGAGTAAGCTC -3’ F2_RGR_PKS12 _1326: 5’-ACGAAACGAGTAAGCTCGTCTGTTCATCGACAAATTCCCGGGTTTTAGA GCTAGAAATAGC-3’

- TracrRNA: endogenous bacterial RNA that links the crRNA to the Cas9 nuclease and can bind any crRNA;

- HDV: Hepatitis Delta Virus Ribozyme.

After the RGR-cassette is transcribed, the guide RNA will be released from the primary transcript automatically due to the presence of the two ribozymes.

The only differences between the two RGR-cassettes were the sequences of the N1-20 _{spacer and, consequently, of the M}6-1_.

M6-1 _GGTCTC Hammer Head ribozime CTGATGAGTCCGTGAGGACGAAACGAGTAAGCTCGTC Spacer N1-20 (1001) GAGACCTTGGTAAGATCCAG Chimeric tracrRNA GTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCGT TATCAACTTGAAAAAGTGGCACCGAGTCGGTGCTTTT Hepatitis Delta

Virus ribozyme GGCCGGCATGGTCCCAGCCTCCTCGCTGGCGCCGGCTGGGCAACATGCTTCGGCATGGCGAATGGGAC

M6-1 _TGAACA Hammer Head ribozime CTGATGAGTCCGTGAGGACGAAACGAGTAAGCTCGTC Spacer N1-20 (1326) TGTTCATCGACAAATTCCCG Chimeric tracrRNA GTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCGT TATCAACTTGAAAAAGTGGCACCGAGTCGGTGCTTTT Hepatitis Delta

Virus ribozyme GGCCGGCATGGTCCCAGCCTCCTCGCTGGCGCCGGCTGGGCAACATGCTTCGGCATGGCGAATGGGAC

The RGR-cassettes were assembled with the PacI linearized pFC322 plasmid containing the Cas9 and the Hygromycin resistance gene, in addition to the AMA1 gene (Figure 4.4).

Figure 4.4: Construction of the plasmid containing the sgRNA and Cas9.

The final Cas9 expressing plasmid contains the Cas9 fused to several NLSs under the control of a constitutive promoter of the Translational Elongation Factor (pTef) and its terminator tTef. The pyrG gene confers Hygromycin resistance in order to select the transformants and to give selective pressure to assure the auto-replication of the plasmid into the fungal cells. Then the oriV control replication plasmid in bacteria and AMP confer ampicillin resistance. The gRNA expression is under the control of the

Glyceraldehide-Phosphato-Dehidrogenase A, a strong and constitutive promoter that is one of the most used promoters for fungi, and the Tryptophan C terminator. Finally the plasmid contains also a truncate AMA1 sequence which is the component that enables the plasmid to replicate autonomously. Figure 4.5 resumes all the Gibson assembly protocol used in the present thesis.

Figure 4.5: Construction of the RGR-cassette and assembly of the Cas9-expression vector.

The new assembled plasmid was directly transferred into E. coli competent cells, thus resulting in a quite large number of bacterial colonies putatively containing the RGR-Cas9-expressing plasmid (1001 and 1326). In order to check the right assembly, a colony PCR was performed on 10 colonies from each of the two assembly protocols. Results from the colony PCR are shown in Figure 4.6, where all colonies (but one in PKS12_1001) resulted to be positive and potentially to be used for fungal transformation.

Figure 4.6: 1% agarose gel showing amplicons obtained from 10 colonies of PKS_1001 (a, lanes A-L) and from 10 colonies of PKS_1326 (b, lanes A-L). Amplicons with primers SB011 and SB012 were 211 bp higher than that obtained from pFC332 (lane ND) used as negative control. CHK= control using water as template.

Five out of the ten clones resulted positive to colony PCR, for each plasmid, were sequenced and sequences were compared with that from pFC332 and from the expected sequence manually constructed. All the five clones for each plasmid resulted to be correctly assembled. Then one plasmid Cas9_PKS12_1001 and one plasmid pCRISPR-Cas9_PKS12_1326 were used for further fungal transformation.

4.2 F. graminearum ITEM 124 transformation

Protoplasts Mediated Transformation (PMT) protocol was used for editing the PKS12 by Cas9 cleavage in F. graminearum ITEM 124. We tested three different Lysing Solutions, differing in the combination and the amount of the lytic enzymes (such as cellulase, driselase, Novozyme...etc). We also tested three different protocols differing in the way that the protoplasts were collected from the Lysing solution. At the end the mix of three different enzymes listed in materials and methods and the collection of protoplasts by filtering in glass wool, allowed to collect an enough number of protoplasts, able to regenerate and ready to be used for the following transformation.

It has been about 30 years since the initial paper on transformation of filamentous fungi was published (Ballance et al., 1983; Tilburn et al., 1983, Yelton et al., 1984). Surprisingly, the actual technique for delivering DNA into the cells has not changed much since then. Most efficient filamentous fungi transformation methods relied on PMT. This is the process whereby protoplasts are made from digested mycelium using a suitable lytic enzyme cocktail, while the fragile digested cells are maintained in a osmotically stabilising buffer (Cantoral et al., 1987). The PMT method is the archetype of fungal transformation, as with many other techniques used for filamentous fungi it originates from Saccharomyces cerevisiae (Hinnen et al., 1978).

Among other methods, Agrobacterium tumefaciens Mediated Transformation (AMT) (de Groot et al., 1998) reaches similar transformation rates as PMT. It involves conjugation with Agrobacterium

tumefaciens and thus requires a completely different preparation process. It

originates from plant infections and uses a bacterium to transfer a T-DNA region of the Ti-plasmid to the genome of the infected plant or fungi.

Despite of the AMT is generally recognized as the most efficient method to transform filamentous fungi, for targeted gene deletion approach such as Cas9 protocol, the preferred method is delivering the system PMT. Whereas only episomal expression of Cas9 protein and a marker gene (Nødvig et al., 2015, Zhang et al., 2015) is needed. This is preferably achieved by using vectors containing an autonomously replicating sequence such as AMA1 from A. nidulans (Aleksenko and Clutterbuck, 1996, 1997), which provides transient expression and extrachromosomal maintenance of the CRISPR-Cas components in the fungal cells.

After the transformation procedure, putatively transformed protoplasts were inoculated on selective medium (PDA with 200 ppm Hygromycin). We performed six different transformations but in only three of them we obtained the regeneration of colonies from protoplasts, as shown in Figure 4.7.

Figure 4.7: F. graminearum ITEM 124 regenerating colonies.

Transformants were generated by using two slightly different protocols: in the first two transformations we inoculated putative transformed protoplasts on non-selective medium (PDA + 1M sorbitol) and after the regeneration of colonies we poured a layer of melted selective medium (PDA 9% + 1M sorbitol + Hygromycin 200 ppm) on the Petri dish. Afterwards, only the colonies that emerged from the selective layer were inoculated in a new Petri dish with selective medium. The second protocols differed form the first one in the way that the putative transformed protoplasts were inoculated directly on a melted selective medium (PDA 9% + 1M sorbitol + Hygromycin 200 ppm) and only the regenerated

colonies were subsequently inoculated in a new Petri dish containing the antibiotic. In both protocols we have inoculated putative transformed colonies on selective medium twice in a row to allow the expression of the Cas9 protein.

After three rounds on PDA without selective antibiotic, six colonies from the first transformation (one for 1001 and five for 1326), eleven from the second one (one from 1001 and ten from 1326) and two from the fifth one (one for 1001 and for 1326) developed and the portion of PKS gene was amplified by PCR, sequenced and compared with the WT. Using Clustal command-line version software (Larkin, 2007) we performed a multi-alignment of putative transformants sequences and WT. We did not observe deletions within the N1-20 _{spacer. As example, the alignment of}

sequences from five PKS_1326 putative transformants with WT is shown in Figure 4.8.

Figure 4.8: example of alignment of the PKS-1326 region from 5 putative transformants with the WT. No deletion occurred within the N1-20 _{spacer (red box)}

The three sub-cultivations on PDA without antibiotic allowed the elimination of the plasmid from fungal cells. Truncated version of AMA1 present in the plasmid should increase the probability of quickly removing the plasmid from the transformants, simply by reducing the selection pressure via consecutive rounds of sub-culturing on non-selective medium.

The AMA1 sequence is a relatively large palindromic structure comprised of a short 0.3kbp central region, which is flanked by two 3kbp inverted MATE1 (Mobile Aspergillus Transformation Enhancers) like elements (Aleksenko and Clutterbuck, 1996).

The Crispr-Cas9 vector equipped with the AMA1 was proficient in self-replicating in A. carbonarius, and this autonomously self-replicating plasmid facilitated the counter selection of the transformation marker. After only three successive rounds of growth on non-selective media, Weyda et al. (2017) saw that their edited strain completely lost its ability to grow in the presence of the selection marker. This gives a big advantage for this transformation technique, as consecutive rounds of transformation using the same marker, and eventually the generation of marker-free mutants can be achieved easily.

Our apparently negative results, due to the lack of transformants, can be due to a low efficiency of the PMT protocol more than to a wrong assembly of the deletion plasmids. According to Shi et al. (2017), the protocol we followed to construct the deletion vector was the most

appropriate for filamentous fungi and the presence of the AMA1 gene within the deletion plasmid guaranteed the elimination of exogenous DNA from the edited strains, thus replying to the main concern connected with the use of genetically modified organisms.

The Crispr-Cas9 technique we used was carried out by using a very large circular plasmid (16.000bp circa), hence, the transformation efficiency can be mainly affected by the uptake of the plasmid by the protoplasts during the transformation. Therefore, several important conditions can be optimized to guarantee promising transformation efficiency, e.g. preparation and storage time of protoplasts, concentration of protoplasts and amount of DNA used for transformation.

Furthermore, to date, there are very few articles where researchers demonstrate that they have succeeded in transforming filamentous fungi with the Crispr-Cas9 technique. In 2017 only 18 papers concerning the use of CRISPR-Cas9 in fungi were available (Figure 4.9) and in 2018 only few ones were added (Zheng et al., 2018; Matsu-ura et al., 2018, Nødvig et al., 2018). Moreover, the low transformation efficiency of this technique is a well-known problem even when it is applied to other organisms (Charpentier E., personal communication).

Presumably this can be caused by two factors:

i) spacer targeting efficiency varies depending on the spacer itself and the targeted gene. Although numerous research groups have developed

software to minimize ON and OFF targeting, it is still difficult to predict the optimal spacer of individual genes. Ideally a spacer sequence would have perfect homology to the target sequence and be unique in the genome. Even so it may not result in the wanted efficiency, as it is now recognized that cleavage efficiency can increase or decrease depending upon nucleotide usage within the selected target sequence. Recent developments have been made by use of machine learning to develop new prediction models based on data from large-scale genome wide CRISPR knockouts. Such models may in time also be applied to predict optimal spacer sequences for activating or repressing genes. However, the efficacy of these spacers maybe influences by other underlying factors that specific for individual promotor regions (Doench et al., 2016).

ii) bulk of Fusarium genus have multicellular or multinuclear hyphae, some of them have two or more genetically diverse nuclei in a shared cytoplasm (heterokaryon). This heterokaryotic nature of these species can affect the overall efficiency of CRISPR-Cas9 systems in filamentous fungi (Katayama et al., 2016). However, there appears to be a mixed correlation between the number of nuclei and the outcome of CRISPR mediated gene knockouts in filamentous fungi. Mutational rates up to 100% were reported in multinuclear Aspergillus oryzae (Katayama et al., 2016) when knocking out the pigment gene yA. While in mononuclear Pyricularia oryzae and

84% and 25–53%, respectively (Arazoe et al., 2015, Fuller et al., 2015). Still, in Aspergillus carbonarius and Aspergillus luchuensis, which are known to contain 2–5 nuclei in each conidium, the effectiveness of CRISPR-Cas9 is perceived to be very low (Nødvig et al., 2015). In any case, it remains a problem that CRISPR-Cas9 works in a nuclei level.

5. Conclusions

In this master thesis we tried to set up a protocol for the genome editing of a Fusarium graminearum mycotoxigenic isolate (ITEM 124) as a proof of concept in order to demonstrate the applicability of the CRISPR-Cas9 technique on this organism.

Currently, fungal genetic modifications can be achieved via many transformation techniques such as Protoplast Mediated Transformation (PMT) and Agrobacterium-Mediated Transformation (AMT). However, low gene targeting frequency remains a main obstacle for precise genome editing in fungi.

Genome editing of filamentous fungi via the CRISPR-Cas9 technology has increased in recent years and it’s considered a potential tool to solve problem of low gene editing frequency in filamentous fungi but it has been established in only a small number of species compared with other organisms such as E. coli, yeast, and zebrafish (Ng et al., 2016, Chung et al., 2017, Gao and Zhao, 2015, Auer et al., 2014). In fact, only few studies investigated the application of genome editing for fungal metabolic engineering, and main works mainly revolved around the feasibility of establishing the CRISPR-Cas9 system in filamentous fungi at all.

Another aspect to keep in mind is that the success of the Crispr-Cas9 technique is closely linked to the fungi DNA repairing systems. There are

two main repairing systems: the NHEJ and the HR, the first one is more active than HR, or HR is less efficient than NHEJ. If this situation is not maintained the DSB induced by Crispr-Cas9 can be repaired correctly and therefore frame-shift events do not happens, resulting in a not-knock-out event.

The use of Cirspr-Cas9 technique opens up new perspectives in the study of molecular biology of fungi. Moreover the use of this technique to silence genes involved in the production of trichothecenes, a virulence factor that determine pathogenicity in fungi such as F. graminearum, can lead to obtaining avirulent strains which could infect the host without producing mycotoxins but, at the same time, can compete for the substrate with mycotoxin-producing wild type strains. In addition, the genome editing of this kind of pathogen can improve the knowledge of the mechanisms controlling the interaction of the pathogen with its host.

From a lab point of view, this technique can also be used to silence individual or multiple genes at the same time of an isolate and study their function in the interaction with antagonistic isolates, such as those belonging to Trichoderma genus. This specific case is very interesting because of the ability of some isolates of Trichoderma (T. gamsii 6085, T.

gamsii 6317 and T. velutinum 3837) to inhibit both fungal growth and DON

production of our isolate of F. graminearum (ITEM 124) as reporter by Matarese and colleagues (2012).

In conclusion Crispr-Cas9 technique exhibited great potential as a tool for genome engineering and investigation of genes functions in many metabolic pathways such as trichothecene production, interaction with hosts and antagonistic species of fungi. Nevertheless, it still needs further studies and enormous efforts to develop an optimized system to obtain better results in filamentous fungi.

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