Regulation and Specificity of the Coprinopsis Cinerea Defensome Against Nematodes

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Doctoral Thesis

Regulation and specificity of the coprinopsis cinerea defensome against nematodes

Author(s): Plaza Gutierrez, David F.

Publication Date: 2014

Permanent Link: https://doi.org/10.3929/ethz-a-010276765

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ETH Library

DISS. ETH NO. 22090

TITLE OF THE DOCTORAL THESIS REGULATION AND SPECIFICITY OF THE COPRINOPSIS CINEREA DEFENSOME AGAINST NEMATODES

A thesis submitted to attain the degree of DOCTOR OF SCIENCES of ETH ZURICH (Dr. sc. ETH Zurich)

presented by DAVID FERNANDO PLAZA GUTIERREZ Master of Science, Uppsala University

born on 02.12.1984 citizen of the Republic of Colombia

accepted on the recommendation of Prof. Dr. Markus Aebi Dr. Markus Künzler Prof. Dr. Adrian Leuchtmann Prof. Dr. Leo Eberl

2014

Acknowledgements

I am very grateful to my supervisors in this project, Prof. Markus Aebi and Dr. Markus Künzler for their precious intellectual input that was crucial for the completion of this thesis. They represent perfect role models to follow for a young and inexperienced scientist like me. I also want to thank the constant presence of my family in the distance, specially my brother

Sebastian and my parents Fernando and Clemencia. I am grateful too to my girlfriend for her unconditional support in the most uncertain times, for showing me the bright side of everything and for putting my feet back to the ground when I needed it most. To my friends in

Switzerland, Juan O., Juan S., Marija, Begonia, Stefan, Gustavo, Elsy and Niels with whom I spend the most joyful non-scientific moments in these four years. I also want to sincerely thank all my scientific mentors, all these adoptive fathers and mothers who guided me through the turbulent waters of science, specially to Clara Spinel and Gabriela Delgado at the

Universidad Nacional de Colombia; Manuel E. Patarroyo and Manuel A. Patarroyo at the

Fundacion Instituto de Inmunologia de Colombia; and Sandra Kleinau at Uppsala University.

"A free man thinks of nothing less than of death; and his wisdom is a meditation not on death but on life" Baruch Spinoza

Table of contents

Summary……………………………………………………………………………………………….i

Zusammenfassung………………………………………………………………………………….iii

Chapter 1………………………………………………………………………………………………1

Introduction: Regulation of gene expression in fungi

Chapter 2………...…………………………………………………………………………………..27

Comparative transcriptomics of the model mushroom Coprinopsis cinerea reveals

tissue-specific armories and a conserved circuitry for sexual development

Chapter 3………...…………………………………………………………………………………..75

Innate immunity in fungi: Transcriptional regulation and target specificity in the

defense response of the model mushroom Coprinopsis cinerea

Chapter 4…………...………………………………………………………………………………111

Cloning, expression and partial functional characterization of putative defense

proteins of Coprinopsis cinerea

Chapter 5…………...………………………………………………………………………………143

Toxicity spectrum analysis of fungal defense lectins in Rhabditid and Diplogasterid

nematodes

Chapter 6……...……………………………………………………………………………………169

Summary, conclusions and outlook

Abstract

In this doctoral thesis, the transcriptional regulation of sexual development and innate defense of Coprinopsis cinerea, a basidiomycetous fungus that grows in horse dung, was studied. In addition, a partial functional characterization of a handful of putative defense proteins from this fungus was performed. Finally, the glycome of some species of Rhabditid and Diplogasterid nematodes was analyzed with regard to the toxic action of an array of fungal defense lectins.

In chapter 1, the state of the art of the transcriptional regulation of gene expression in fungi is reviewed. The review includes the different mechanisms used by fungi to monitor the biotic and abiotic components of the environment they inhabit, as well as the signaling pathways leading to changes in their gene expression. Gene expression programs associated to fungal development and their regulatory connection to secondary metabolism are explained. Finally, the role of post-transcriptional regulation and the histone code in the control of gene expression are discussed.

Chapter 2 presents the developmental transcriptome of C. cinerea A43mutB43mut comparing stage 1 primordia and vegetative mycelium. Differential expression analysis shows that, on the one hand, stage 1 primordia expresses loci encoding nematotoxic/insecticidal lectins and protease inhibitors, whereas vegetative mycelium mainly expresses genes encoding putative bactericidal proteins. These defense proteins are thought to match the predators and competitors confronted by the two different tissues in the environments where these tissues develop.

Transcriptional regulation of the innate defense of C. cinerea vegetative mycelium against bacteria and fungivorous nematodes is shown in chapter 3. Three nematotoxic lectins were found to be significantly induced in C. cinerea by Aphelenchus avenae predation. Moreover, expression of loci encoding putative lysozymes was found to be increased upon growing C. cinerea in the presence of Escherichia coli and Bacillus subtilis. Lack of induction of these loci

i following mechanical injury of the hyphae or co-culture with the bacterivore Caenorhabditis elegans shows that C. cinerea can recognize different environmental stimuli and trigger a

"stimulus-specific" transcriptional response. Lastly, antisense transcription was identified for a major fraction of the protein-coding genome; the proportion of antisense transcription was found to significantly change under some culture conditions for a handful of open reading frames.

A partial functional characterization of four putative defense protein from C. cinerea is described in chapter 4. First, C. cinerea protease inhibitor A 1 (CCIA1) was shown to be s

Subtilisin-inhibitor that is toxic to C. elegans L4 larvae. Second, the vegetative mycelium- specific Ricin B domain-containing protein CC1G_05299 was shown to not bind lactose and to not be toxic to C. elegans and Aedes aegypti. Lastly, two additional putative defense proteins were shown to be insoluble when produced recombinantly in E. coli or Pichia pastoris.

In chapter 5, the toxicity spectrum of several fungal defense lectins against different species of Rhabditid and Diplogasterid nematodes was explored. Lectins targeting Gal-GalNAc motifs, commonly found in glycolipids, were shown to be toxic to all the species tested. On the other hand, large species-specificity differences were observed for fucose-binding lectins. Different mechanisms for the appearance of resistance to lectin toxicity in nematodes are discussed.

These findings reflect part of the complexity behind the interaction between fungi and the nematodes that predate on them. On the one hand, they show that fungi evolved an innate defense system comparable in specificity and inducibility to the animal innate immune system.

On the other hand, they demonstrate that nematodes have a versatile and diverse glycorepertoire that allows them to naturally generate resistance to fungal defense lectins.

Zusammenfassung

In der vorliegenden Doktorarbeit wurde die transkriptionelle Genegulation des Modellhutpilzes

Coprinopsis cinerea während der sexuellen Entwicklung sowie der angeborenen Abwehr gegenüber Bakterien und pilzfressenden Nematoden untersucht. Einige putative

Abwehrproteine, die im Zuge dieser Studie identifiziert wurden, konnten zusätzlich auf funktioneller Ebene charakterisiert werden. Ferner wurde das Glykom einiger Rhabditid und

Diplogasterid Nemaatodenspezies im Hinblick auf deren Sensitivität gegenüber einer Auswahl toxischer pilzlicher Abwehrproteine analysiert.

Kapitel 1 gibt einen Überblick über die transkriptionelle Regulation der Genexpression in Pilzen im Allgemeinen. Es werden unterschiedliche Mechanismen beleuchtet, die es Pilzen erlauben, biotische, wie auch abiotische Komponenten ihrer Umgebung wahrzunehmen und als

Reaktion auf diese Signale mittels Signaltransduktionswegen schliesslich Änderungen im

Expressionsprofil des Genoms herbeizuführen. Im Kapitel werden zudem die Rolle post- transkriptioneller Regulation und des Histone Codes besprochen.

Kapitel 2 beinhaltet die differenzielle Transkriptomanalyse während der sexuellen Entwicklung in C. cinerea A43mutB43mut. Dabei wurden Stadium 1 Primordien mit vegetativem Myzel verglichen. Die Analyse der differenziellen Geneexpression von zwischen Stadium 1

Primordien und vegetativem Myzel zeigt, dass in Stadium 1 Primordien Genloci aktiviert werden, die für nemato-und entomotoxische Lektine und Protease-Inhibitoren kodieren; in vegetativem Myzel hingegen werden hauptsächlich putative antibakterielle Proteine exprimiert. Diese differenzielle Expression verschiedener Abwehrproteine spiegelt möglicherweise die unterschiedlichen Prädatoren und Konkurrenten der beiden Gewebe wider, mit denen sie im jeweiligen Habitat konfrontiert werden.

Die transkriptionelle Regulation der angeboren Abwehr in vegetativen Myzel C. cinerea gegen

Bakterien, sowie gegen fungivore Nematoden, wird in Kapitel 3 behandelt. In Anwesenheit von

iii

Escherichia coli oder Bacillus subtilis werden in C. cinerea vermehrt Lysozm kodierende

Genloci exprimiert. Wohingegen drei nematotoxische Lektine indentifiziert werden konnten, die durch Prädation von Aphelenchus avenae induziert werden. Dass keiner dieser Genloci durch mechanische Destruktion des vegetativen Myzels oder durch Co-Kultivierung mit dem bakteriovoren Nematode C. elegans induziert werden konnte, impliziert, dass C. cinerea differenziert, mit einer für Prädator- oder Konkurrent-spezifischen Änderung in der

Genexpression, auf unterschiedliche Stimuli reagieren kann. Für viele Protein-kodierende

Gene konnte zudem 'Antisense'-Transkription identifiziert werden, die sich, abhängig von

Kulturbedingungen, signifikant für einige offene Leseraster ändern kann.

Kapitel 4 beschreibt die funktionelle und biochemische Charakteriesierung vier in dieser

Studie identifizierte Abwehrproteine. Es konnte gezeigt werden, dass der C. cinerea Protease

Inhibitor CCIA1 ein Subtilisin-Typ Protease Inhibitor ist, der toxische Wirkung auf C. elegans

L4 Larven aufweist. Das Ricin-Domänen-Protein (CC1G_05299), das spezifisch im Myzel exprimiert wird, zeigte keine toxische Wirkung, weder gegen C. elegans noch gegenüber

Aedes aegypti. Zudem konnte keine Lactose Bindung des Proteins festgestellt werden. Zwei weitere Abwehrproteine konnten in E.coli sowie in Pichia pastoris nur in unlöslicher Form produziert werden.

In Kapitel 5 wurde das Toxizitätsprofil mehrerer pilzlicher Lektine gegen einige Rhabditid und

Diplogastrid Spezies untersucht. Lektine mit einer Spezifität für Gal-GalNAc, ein gängiges

Motiv auf Glycosphingolipiden, waren toxisch für alle getesteten Spezies. Lektine mit einer

Fucose-Spezifität, zeigten hingegen unterschiedliche Toxizität gegenüber den getesteten

Spezies. Potenzielle Mechanismen, die zu Resistenzbildung in diesen Spezies führen könnten, werden zusätzlich diskutiert.

Die Ergebnisse reflektieren einen Teil der Komplexität der antagonistischen Interaktion zwischen einem Hutpilz und pilzfressendenen Nematoden. Sie zeigen, dass der Pilz eine

Protein-basierte, angeborene Abwehr besitzt, die in ihrer Spezifität und Induzierbarkeit der tierischen ähnlich ist. Andererseits zeigen sie auch, dass Nematoden ein vielfältiges

Glycorepertoire besitzen, das eine natürliche Resistenz gegenüber pilzlichen Lektinen darstellt.

Chapter 1 Introduction: Regulation of gene expression in fungi

Chapter 1

Introduction:

Regulation of gene expression in fungi

David Fernando Plaza1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

Chapter 1 Introduction: Regulation of gene expression in fungi

Regulation of gene expression in fungi

David Fernando Plaza1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

Abstract

Several layers of regulation determine how much a gene is expressed at a given condition. A lot of what is currently known about gene regulation in fungi is derived from pioneering studies in the yeast Saccharomyces cerevisiae; only recently, new genome sequencing technologies and genome manipulation tools have allowed us to elucidate the gene regulation of filamentous fungi. Fungi modulate their gene expression according to environmental conditions in order to adapt to their ecological niche. They use receptors and downstream signaling cascades to modulate the expression of specific genes. At least 37 classes of transcription factors have been described from the fungal genome sequences available. Among these, the Velvet family of transcription factors is of particular interest because their members structurally resemble regulatory proteins involved in development and defense of animals and plants, suggesting that some ancient gene regulatory mechanisms are conserved in all eukaryotes. This short review provides a general glance on the different strategies fungi use to sense and respond to their environment, and discusses the ways by which fungi adjust their gene expression to stress, microbial interactions and development. In addition, current knowledge on the importance of epigenetics and antisense- mediated regulation of gene expression in fungi are briefly discussed.

Keywords: abiotic stress, biotic stress, fungal development, secondary metabolism, environment sensing, signaling pathways, transcription factors, histone code, posttranscriptional regulation

Chapter 1 Introduction: Regulation of gene expression in fungi

Regulation of fungal gene expression in response to abiotic stress

Organisms permanently interact with their environment and adapt the expression of their genome to the challenges they are confronted with. These challenges can be abiotic or biotic. Different studies have presented evidence that fungi can trigger compensatory mechanisms to respond to environmental stresses. For example, the pathogenic ascomycete

Paracoccidioides brasiliensis expresses genes involved in the maintenance of redox homeostasis as well as genes involved in cell wall remodeling in response to oxidative stress

[1]. Similarly, the arbuscular mycorrhizal fungus Glomus intraradices turns on the expression of the aquaporin GintAQP1 when subjected to osmotic stress [2].

Enjalbert and collaborators have shown that Candida albicans, an opportunistic pathogen of humans, expresses different genes in response to different stresses [3]. For instance, C. albicans specifically increases the transcription of genes encoding the heat shock proteins

HSP12, HSP70, HSP78, and HSP104 which are involved in the structural protection of protein stability when the growth temperature is switched from 23 to 37°C [3]. In addition, C. albicans also induces the specific expression of a glutathion reductase, a thioredoxin and a thioredoxin oxidase when exposed to oxidative shock [3].

Regulation of fungal gene expression in response to biotic stress

Aspergillus nidulans induces the expression of a gene cluster associated to polyketide synthesis when it is grown in close physical contact with the bacterium Streptomyces rapamycinicus (previously called Streptomyces hygroscopicus). This cluster was found to be responsible of the synthesis of orsellinic acid, lecanoric acid and the cathepsin K inhibitors F-

9775A and F-9775B [4]. Intriguingly, genes involved in stress response, carbohydrate metabolism and cellular transport were shown to be down-regulated when the ascomycete

Magnaporthe oryzae was grown with pathogenic Lysobacter enzymogenes C3. These genes were not modulated when M. oryzae interacted with the non-pathogenic strain L. enzymogenes DCA, indicating that pathogenic bacteria can repress the expression of

Chapter 1 Introduction: Regulation of gene expression in fungi defense-related genes in fungi [5]. As a form of defense against antifungal metabolites, the phytopathogenic ascomycete Botrytis cinerea induces the expression of ATP-binding cassette (ABC)-containing transporters in response to phloroglucinols and phenazines produced by some Pseudomonas isolates [6-8].

Saupe and collaborators described a novel prion-mediated mechanism of signal detection in the context of heterokaryon incompatibility (HI) in Podospora anserina [9]. HI is an apoptosis- like reaction that takes place when two genetically incompatible hyphae of the same species fuse [10]. HI is mediated by the binding of cytoplasmic HET domain-containing proteins from the two fusing hyphae. Het-s is a prion-like HET domain-containing protein that forms fibrilar polymers in Podospora. Het-s+, on the contrary, is a soluble folding variant of Het-s. Fusion of a Het-s and a Het-s+ strain results in the conversion of the Het-s+ into Het-s without triggering a cell death response. In contrast, fusion of Het-s and Het-S, an incompatible isoform of the protein, leads to cell death in proximity to the fusion region. The exact cause of toxicity is still unknown but it's thought to involve the formation of Het-s/Het-S heterodimers

[9].

Coordination of fungal development and secondary metabolism

When filamentous fungi undergo sexual reproduction, monokaryotic hyphae from two individuals of compatible mating type fuse (plasmogamy) to form dikaryotic hyphae where every cell contains two different types of nuclei derived from the two respective parental individuals [11]. Thereafter, environmental factors such as nutrient depletion or light, trigger the fusion of nuclei and the start of meiosis. The induction of sexual reproduction in filamentous fungi goes usually along with extensive cellular differentiation and the formation of specialized fruiting bodies for spore dispersal [12-14].

Gene expression associated with sexual development has been studied in both ascomycetes and basidiomycetes. The results of such studies show that there are few commonalities in

Chapter 1 Introduction: Regulation of gene expression in fungi the general expression programs between different species. Antisense transcripts mapping to 281 genes and up-regulation of several orphan loci were observed in the ascomycetous fungus Pyronema confluens. Comparative transcriptome analysis with and complementation experiments in Sordaria macrospora showed that the function of the transcription factor

Pro44 in the regulation of sexual development is conserved in ascomycetes [15]. On the other hand, the fruiting body transcriptome from the Witches' Broom Disease-causing basidiomycetous fungus Moniliophthora perniciosa revealed that genes encoding hydrophobins, Rho-GEFs, extensin precursors, glucose transporters and cytochrome p450 monooxygenases are induced during sexual development [16]. Hydrophobins and loci encoding proteins involved in intracellular trafficking and stress responses were found to be up-regulated in primordia from the Shiitake mushroom Lentinula edodes [17]. Transcriptome analysis in Schizophyllum commune revealed that, besides hydrophobins, one third of all the transcription factors encoded in the genome is differentially expressed during sexual development. One of these differentially expressed transcription factors, Fst4, was shown to be necessary for the induction of fruiting body formation [18]. Finally, genes encoding small secreted proteins and loci involved in cell wall remodeling were shown to be up-regulated in fruiting bodies of the ectomycorrhizal fungus Laccaria bicolor [19].

Loci encoding enzymes responsible for the biosynthesis of secondary metabolites in fungi can be found clustered in close proximity in the genome [20, 21]; furthermore, some of these genes have been shown to be transcriptionally co-regulated (Figure 3) [20]. In the ascomycete Fusarium graminearum, clusters of genes in close chromosomal proximity can be co-regulated at the transcriptional level [22].

Chapter 1 Introduction: Regulation of gene expression in fungi

Figure 3. Transcriptional co-regulation of enzymes involved in secondary metabolite production. (A) The gene cluster encoding enzymes responsible of the biosynthesis of

Sterigmatocystin is turned on when A. nidulans is grown on solid medium. Every line represents one of the genes in the cluster. The X-axis corresponds to independent experiments, 36 in liquid medium and 8 in solid medium. (B) Polyketide synthesis gene cluster encoding the enzymatic machinery involved in the synthesis of Sterigmatocystin.

Figure adapted from [20].

Intriguingly, conditions triggering the production of fungal secondary metabolites coincide with those necessary for the initiation of sexual or asexual development in Aspegillus

(nutrient availability, pH, temperature) [23]. Development-associated secondary metabolites can be divided in two categories: those which are essential for spore production, such as some linoleic acid derivatives [24], and those which are dispensable (e. g. aflatoxin). 13S- hydroperoxylinoleic acid was shown to induce conidia production when applied exogenously to A. flavus [24], demonstrating that products of the secondary metabolism can have a direct effect in the initiation of asexual development of ascomycetes. In addition, linoleic acid derivatives such as behenic and sebacic acid, have been shown to induce production of the carcinogenic natural product aflatoxin [25]. Furthermore, mutations affecting conidia formation in A. parasiticus impair aflatoxin biosynthesis [26]. However, aflatoxin-deficient strains carrying a mutated version of AflR, a transcription factor specifically regulating 6

Chapter 1 Introduction: Regulation of gene expression in fungi sterigmatocystin/aflatoxin gene clusters, are capable of producing spores and sclerotia [23,

27], indicating that production of aflatoxin is subordinated to the initiation of asexual development.

Signals, receptors and signal transduction pathways

Generally, the external stimuli are perceived by receptors in the plasma membrane and transformed into changes in gene expression by signaling cascades which are conserved among phylogenetically distant eukaryotes [28]. G-protein coupled receptors (GPCRs) are proteins anchored to the cell by 7 membrane-spanning helices and a fundamental component of the environment sensing and transduction machinery in fungi [29]. Fungal

GPCRs can be classified into six classes based on their function and sequence similarity:

Ste2-like pheromone receptors, Ste3-like pheromone receptors, carbon/amino acid receptors, putative nutrient receptors, cAMP receptor-like proteins and microbial opsins [29].

Pheromone emission and sensing are crucial to attract compatible mating partners. The

GPCRs GprA and GprB have been described in A. nidulans as responsible for pheromone sensing during self-fertilization [30]. GPCRs are also involved in the sensing of nutrient availability. In the ascomycete Neurospora crassa, the quality of the carbon source available in the medium is sensed by the GPCR Gpr-4. In a medium were glycerol is the only carbon source, Gpr-4 forms a complex with the cytoplasmic Gα protein GNA-1 which leads to the activation of the adenylyl cyclase CR-1 and the production of cAMP upon glycerol binding to

Gpr-4 [31].

Some membrane transporters play an additional role in sensing the chemical composition of the environment. In Ustilago maydis, for example, the ammonium transporter Ump2 was shown to sense the concentration of ammonium in the medium [32]. S. cerevisiae defective in the methylammonium permease Mep2 is unable to form pseudohyphae under low- ammonium conditions [33]; however, heterologous expression of Ump2 restores pseudohyphae formation in Saccharomyces [32].

Chapter 1 Introduction: Regulation of gene expression in fungi

Similarly, light is a crucial developmental cue for some fungi [34, 35]. Photosensory proteins have been identified but little is known about the downstream events following light sensing

[36]. Opsins are structurally conserved membrane-anchored proteins carrying a chromophore coupled to a conserved lysine residue. The photosensory opsin white collar-1

(WC-1) is part of the circadian clock of N. crassa. WC-1 contains a photosensitive flavin- interacting domain and a zinc-finger domain which allows it to interact directly with DNA. In the dark, WC-1 forms a heterodimer with the DNA-binding protein WC-2. When exposed to light, WC-1 releases WC-2 and it is thought to bind a second yet unidentified molecule [37].

It has been shown that the MAP kinase and the cAMP-PKA pathways are involved in growth, development and virulence in filamentous fungi [38]. These two pathways allow these organisms to respond to pheromones, osmolarity, nutrient conditions or different host-derived signals [38] such as surface hydrophobicity [39]. MAP kinase (MAPK) signaling has been shown to regulate mating in the yeasts S. cerevisiae and Schizosaccharomyces pombe. For example, orthologs to MAPKKK (Byr2/Ste8), MAPKK (Byr1/Ste1) and MAPK (Spk1) have been mutated and shown to be required for sporulation and conjugation in these two yeasts

[40]. A similar role of MAP kinase signaling in reproduction has been reported for other fungi such as Ustilago maydis [41], C. neoformans [42] and N. crassa [43]. Additionally, MAP kinase cascades have been demonstrated to mediate appresorium formation and virulence in the rice blast fungus (ascomycete) M. grisea [44, 45].

Signal transduction mediated by production of cAMP and activation of PKA is important for pathogenicity in the ascomycete Colletotrichum orbiculare. In C. orbiculare, RPK1 and CAC1 are homologs to a regulatory subunit of PKA and an adenylate cyclase, respectively.

Mutations in these two genes render C. orbiculare avirulent to cucumber plants [46]. In the same way, Cryptococcus neoformans carrying a mutation in the PKA PKR1, was observed to be nonpathogenic, failed to produce melanin or capsule and was sterile [47].

Chapter 1 Introduction: Regulation of gene expression in fungi

In the cell, concentration of Ca++ is higher extracellularly and in the lumen of the endoplasmic reticulum (ER) than in the cytoplasm. However, Ca++ gains access to the cytoplasm upon activation of Ca++ channels in the cytoplasmic membrane and the membrane of the ER.

++ These Ca channels open in response to inositol triphosphate (IP3)-binding which in turn is produced by the cleavage of phosphatidylinositol 4,5-bisphosphate (PIP2) into diacyl glycerol

++ (DAG) and IP3 by the membrane-associated enzyme Phospholipase C. Released Ca can thus bind calmodulin or protein kinase C (PKC). Upon Ca++ binding, calmodulin releases the protein phosphatase calcineurin which can then activate different transcription factors [48]. In

C. albicans calmodulin inhibition leads to the inability of transitioning from yeast to filamentous growth [49]. Moreover, C. albicans mutants lacking calcineurin subunit A are not virulent in the mouse model [50]. In C. neoformans, melanin is a virulence factor [51] and

DAG is a component of the melanin synthesis pathway. Cell-wall homeostasis is partially mediated by the proper activation of PKC by DAG. A mutation in the DAG-binding domain of

PKC results in the improper association of laccase to the cell membrane and a significant reduction in the amount of melanin produced [52].

Transcription factors

Thirty seven classes of transcription factors have been characterized in fungi and 6 of them are specific to this kingdom [53]. Todd and collaborators recently published a systematic analysis on the presence and abundance of these 37 classes in at least 77 ascomycetes and

31 basidiomycetes [54], accounting for 36636 genes in the species studied (Figure 1). They showed that there are classes which are unique to either basidiomycetes (C2H2 and CCHC zinc-finger) or ascomycetes (Zn2Cys6 and PF04082), suggesting that the gene regulatory networks of the two fungal phyla were subject to a major differentiation after their evolutionary divergence [54].

Chapter 1 Introduction: Regulation of gene expression in fungi

Figure 1. Prevalence of different PFAM transcription factor classes in basidiomycetous and ascomycetous species. Colors indicate the number of loci corresponding to a specific class in each of the fungal species analyzed. Red represents a high number of members in a class, whereas green indicates a low number of members.

Hierarchical clustering analysis shows that phylogenetically related species clustered together according to the transcription factor classes and the abundance of loci corresponding to these classes encoded in their genomes. Figure adapted from [54].

Combinatorial binding of transcription factors to promoter regions regulates gene expression in fungi. In the yeasts S. cerevisiae, Kluyveromyces lactis and C. albicans, it has been shown that transcription factor binding sites can be lost or gained in promoter regions during the evolution of orthologous loci, altering the conditions under which a gene is turned on or off,

Chapter 1 Introduction: Regulation of gene expression in fungi which largely depend on the combination of transcription factors being capable of binding to a particular promoter [55].

Interestingly, the up-regulation of Velvet domain-containing proteins is conserved during sexual development in the basidiomycetes C. cinerea (Plaza et al, 2014), L. bicolor [19] and

S. commune [18]. Velvet-mediated regulation of gene expression was first shown to link reproduction and secondary metabolite production is ascomycetes [56]. Intriguingly, the

Velvet-domain of VosA from A. nidulans structurally resembles the Rel domain of NF-κB, the master regulator of the innate immune response in vertebrates (Figure 2A), suggesting that this fold is highly conserved structurally and functionally during eukaryote evolution. Similar to NF-κB, VosA was also shown to form VosA-VosA homodimers or VosA-VelB heterodimers that could potentially bind different gene regulatory sequences [57]. ChIP-seq experiments with the Velvet domain-containing proteins VosA and Ryp revealed that these two proteins bind different DNA motifs in the promoters flanking the genes that they regulate (Figure 2B).

Chapter 1 Introduction: Regulation of gene expression in fungi

Figure 2. Velvet domain-containing proteins are trans-regulatory elements of gene expression that recognize diverse promoter motifs in the loci they control. (A)

Superposition of VosA Velvet and NF-κB Rel-N domains showing high structural similarity.

Protein-DNA interphase from the two superposed domains is shown in the right hand pane.

(B) VosA and (C) Ryp recognition motifs as deduced from ChIP-on-chip. Figure adapted from [57, 58].

Histone modifications

Epigenetic marks on histones, such as methyl or acetyl groups, represent another layer of regulation in the gene expression of fungi. The recent sequencing of hundreds of fungal genomes [59] has revealed the presence of several putative histone methyltransferases, demethylases, acetyl transferases and deacetylases capable of modifying lysine, serine or arginine residues on histones, thus silencing or activating different chromatin regions by

Chapter 1 Introduction: Regulation of gene expression in fungi altering the extent of their packing and accessibility to transcription factors and RNA polymerases [60, 61]. This post-translational modification of histones has been extensively studied in the model ascomycete A. nidulans, showing that at least five lysines, one serine and one arginine at the N-terminus tails of the histones H3 and H4 are modified by a battery of enzymes (Figure 4) [61]. Recent evidence has shown that LaeA, an A. nidulans methyl transferase, might regulate secondary metabolite production by altering the methylation state of nucleosomes containing gene clusters involved in the biosynthesis of metabolites such as sterigmatocystin, penicillin, and lovastatin [62, 63].

Figure 4. N-terminus modifications on histones 3 (H3) and 4 (H4) of A. nidulans.

Histone post-translational modifications described for A. nidulans are shown. AC: acetylation;

ME: methylation; PH: phosphorylation. Figure adapted from [61].

A. clavatus [64] and A. oryzae [65], have been shown to use histone post-translational modifications to regulate the expression of genes involved in secondary metabolism. The expression of a secondary metabolite gene cluster in A. nidulans upon interaction with S. rapamycinicus was shown to be epigenetically regulated by Saga/Ada-mediated acetylation of H3K9 [66]. On the contrary, A. oryzae histone de-acetylases hdaA/Aohda1, hdaB/Aorpd3, hdaD/Aohos2 and hst4/AohstD were shown to also play a role in responding to osmotic and oxidative stress, and controlling cell wall homeostasis and chromatin integrity [67]. A similar 13

Chapter 1 Introduction: Regulation of gene expression in fungi role in the response to oxidative stress has been suggested for the arginine methyltransferases RmtA and RmtC from A. nidulans [68].

Post-transcriptional regulation and antisense transcription

RNA interference is thought to have evolved as a defense mechanism against viruses and transposable elements in eukaryotes [69]. The minimal enzymatic machinery necessary for

RNA silencing comprises at least one RNA-dependent RNA polymerase, one Dicer, one

Argonaute polypeptide and one Piwi-like protein [70]. Phylogenetic analysis on the presence or absence of this machinery in different clades has led to the conclusion that the last eukaryotic common ancestor (LECA) was capable of post-transcriptionally silencing gene expression [71]. Accordingly, pathways of RNA interference have been characterized in fungi

[72]. Two Dicer orthologs were identified in the ascomycete M. oryzae; nevertheless, only one of them (MDL-2) was shown to be responsible of hairpin-dependent RNA silencing in this fungus [73]. During evolution, S. cerevisiae lost the capacity of post-transcriptionally regulating gene expression via RNA interference. Remarkably, heterologous expression of

Dicer and Argonaute proteins from S. catellii in S. cerevisiae restored its ability to silence endogenous retrotransposons [74]. In N. crassa, introduction of DNA sequences with homology to endogenous genes results in their silencing in an Argonaute-dependent phenomenon called quelling [35, 75]. At least 25 miRNA-like sRNAs (milRNAs) have also been discovered in N. crassa. These milRNAs post-trancriptionally regulate gene expression in a Dicer-independent manner [76].

Natural antisense transcription has been recently studied in several species of ascomycetes and basidiomycetes [77]. Three mechanisms by which natural antisense transcription can control gene expression in fungi have been characterized (Figure 5) [77]. Firstly, transcriptional interference is caused by the direct collision of RNA polymerases transcribing in close proximity over two complementary strands of DNA [77]. Secondly, antisense transcription can recruit chromatin-remodeling enzymes to a locus, resulting in its silencing or

Chapter 1 Introduction: Regulation of gene expression in fungi activation [77]. Finally, antisense-mediated intron retention is caused by the inability of the splicing machinery to remove introns from double-stranded RNA [77]. In fungi, evidence shows that antisense transcription partially regulates gene expression associated to mating and meiosis [78], phosphate [79] and carbon [80] metabolism, aging [81, 82], circadian clock

[83] and pathogenesis [84].

Figure 5. Mechanisms of gene expression regulation mediated by natural antisense transcription. (A) Transcriptional interference. (B) Chromatin remodeling mediated by antisense transcription. Histone modifications are shown as colored vertical boxes. HDAC:

Histone deacetylase; HMT: Histone methyltransferase (C) Antisense-mediated intron retention. Scenarios in the presence or absence of natural antisense transcripts for each mechanism illustrated are shown. Figure adapted from [77].

Conclusions

Fungi use several regulatory check points to control which genes are expressed and to which extent. Environmental changes are sensed by receptors that then trigger conserved signaling cascades that end up in the activation of at least 37 different classes of regulatory proteins that gain access to the nucleus and activate or repress gene transcription. Sexual development and secondary metabolite production have been shown to be interconnected by

Chapter 1 Introduction: Regulation of gene expression in fungi a common set of transcription factors in ascomycetes. Some of these regulatory proteins are conserved in basidiomycetes at the sequence and transcriptional levels and might also play a role in their sexual development. Experimental evidence has also shown that fungi can regulate their gene expression in response to specific environmental stresses or microbial interaction partners. Finally, the role of epigenetic marks in the form of methyl or acetyl groups on specific histone residues, as well as the relevance of antisense transcription in the activation or modulation of gene expression in fungi, have been experimentally demonstrated.

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Chapter 2 Developmental transcriptomics of C. cinerea

Chapter 2

Comparative transcriptomics of the model mushroom Coprinopsis

cinerea reveals tissue-specific armories and a conserved circuitry

for sexual development

David Fernando Plaza1, Chia-Wei Lin1, Niels Sebastiaan Johannes van der Velden1, Markus

Aebi1, Markus Künzler1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

Contributions:

Sample preparation

RNA extraction and quality control

Differential expression analysis

Comparative transcriptomics analysis

qRT-PCR validation of gene expression

Published in BMC Genomics. 2014 Jun 19; 15(1): 492.

Chapter 2 Developmental transcriptomics of C. cinerea

Comparative transcriptomics of the model mushroom Coprinopsis

cinerea reveals tissue-specific armories and a conserved circuitry

for sexual development

David Fernando Plaza1, Chia-wei Lin1, Niels Sebastiaan Johannes van der Velden1,

Markus Aebi1, Markus Künzler1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Zürich, Switzerland

Abstract

It is well known that mushrooms produce defense proteins and secondary metabolites

against predators and competitors; however, less is known about the correlation between

the tissue-specific expression and the target organism (antagonist) specificity of these

molecules. In addition, conserved transcriptional circuitries involved in developing sexual

organs in fungi are not characterized, despite the growing number of gene expression

datasets available from reproductive and vegetative tissue. The aims of this study were: first,

to evaluate the tissue specificity of defense gene expression in the model mushroom

Coprinopsis cinerea and, second, to assess the degree of conservation in transcriptional

regulation during sexual development in basidiomycetes. In order to characterize the

regulation in the expression of defense loci and the transcriptional circuitries controlling

sexual reproduction in basidiomycetes, we sequenced the poly(A)-positive transcriptome of

stage 1 primordia and vegetative mycelium of C. cinerea A43mutB43mut. Our data show

that many genes encoding predicted and already characterized defense proteins are

differentially expressed in these tissues. The predicted specificity of these proteins with

regard to target organisms suggests that their expression pattern correlates with the type of

antagonists these tissues are confronted with. Accordingly, we show that the stage 1

primordium-specific protein CC1G_11805 is toxic to insects and nematodes. Comparison of

our data to analogous data from Laccaria bicolor and Schizophyllum commune revealed that

Chapter 2 Developmental transcriptomics of C. cinerea

the transcriptional regulation of nearly 70 loci is conserved and probably subjected to stabilizing selection. A Velvet domain-containing protein was found to be up-regulated in all three fungi, providing preliminary evidence of a possible role of the Velvet protein family in

sexual development of basidiomycetes. The PBS-soluble proteome of C. cinerea primordia

and mycelium was analyzed by shotgun LC-MS. This proteome data confirmed the presence

of intracellular defense proteins in primordia. This study shows that the exposure of different

tissues in fungi to different types of antagonists shapes the expression pattern of defense loci in a tissue-specific manner. Furthermore, we identify a transcriptional circuitry conserved among basidiomycetes during fruiting body formation that involves, amongst other transcription factors, the up-regulation of a Velvet domain-containing protein.

Keywords: Fungal defense; sexual development; C. cinerea; primordia; RNA-seq; comparative transcriptomics; tissue-specific LC-MS proteome.

Chapter 2 Developmental transcriptomics of C. cinerea

Introduction

The last eukaryotic common ancestor (LECA) was facultatively sexual and evolved nearly

1.5 billion years ago in the Proterozoic eon [1]. Sexual reproduction (SR) shares common features across the eukaryotic lineage such as ploidy changes, meiotic recombination and cell-cell recognition between gametes followed by cellular fusion and zygote formation [2].

An increase in the genetic diversity of the population, making it more adaptable to changing

environmental conditions, as well as the dilution of deleterious mutations out of the gene pool are the most remarkable evolutionary innovations achieved by SR [3]. Despite all these clear benefits, sex is energetically expensive and entails a higher chance of genetic and organelle conflicts [2].

During sexual reproduction, fungi undergo dramatic morphological changes driven by environmental conditions such as light, nutrient availability and grazing by predators [4]. In basidiomycetes, mushroom development starts with intense localized hyphal branching leading to the formation of hyphal knots. These branching hyphae further aggregate to form

1-2 mm secondary nodules where cell differentiation leads to the establishment of bipolar

primordia containing all the tissues observed in the mature fruiting body [5]. As a last step,

primordia develop to mature fruiting bodies mainly by cellular expansion [6]. Due to their

hyphal density, primordia and fruiting bodies are attractive to predators including mollusks,

arthropods and nematodes [4].

Coprinopsis cinerea has been used as a model basidiomycete since the mid-1950s [7] due

to its saprobic lifestyle, its rapid growth and the feasibility of producing fruiting bodies under

defined laboratory conditions [6]. In nature, C. cinerea grows on horse dung [6], a eutrophic

substrate rich in competing microorganisms, such as Firmicutes, Bacteroidetes and

Proteobacteria [8]. The recent sequencing of the C. cinerea genome [9] allows the study of

Chapter 2 Developmental transcriptomics of C. cinerea this organism gene expression on transcriptome and proteome level at different developmental stages or under a variety of environmental settings.

Morphological changes and environmental signals during fruiting body formation in C. cinerea are well described [6]; nonetheless, comparably little is known about the molecular machinery driving sexual reproduction processes in this basidiomycete. Recently, mutations blocking fruiting body development at different stages or altering mushroom morphology were identified [10-15]. In addition, dst2 and dst1, encoding a blue-light photoreceptor and a flavin adenine dinucleotide-binding protein, were shown to play a role in blue light sensing. In agreement with previous experiments, strains defective in these two proteins were unable to form fruiting bodies, showing that blue light is an essential environmental trigger of mushroom development [16]. Most recently, C. cinerea strains carrying mutations in the putative component of the SWI/SNF chromatin remodeling complex snf5 (CC1G_15539) were shown to be defective in fruiting initiation, suggesting that epigenetic reprogramming of loci occurs during fruiting body formation [17].

Aerial fruiting bodies are an attractive prey for predators and thus are protected by a battery of defense molecules (toxins) including proteins [18-21], peptides [22] and secondary metabolites [23-25]. Some of these toxins are known to be specifically produced in the fruiting body and not in the vegetative mycelium [26, 27]. For instance, cytoplasmic lectins showing a broad range of non-self-carbohydrate specificities, also referred to as fruiting body lectins due to their specific expression pattern, have been shown to exert toxicity to nematodes, insect larvae and amoeba [18]. Protein-mediated inhibition of serine proteases

[20], proteolytic degradation of predator-derived proteins [28, 29] and sequestration of biotin

[19] are other strategies of basidiomycetous fruiting bodies to dissuade predators.

Vegetative mycelium, in contrast, digests extracellular carbon macromolecules in the growth substrate, such as cellulose and lignin, into smaller degradation products which are absorbed by the growing hyphae. At the same time, these smaller molecules become

Chapter 2 Developmental transcriptomics of C. cinerea

available for competing bacteria that profit from the fungal enzymatic machinery [30]. As a response to bacterial competitors, fungi have evolved secreted antimicrobial proteins.

Mygind and colleagues presented recent evidence that vegetative mycelia of fungi secrete cysteine-stabilized antibacterial peptides which play a role in the arms race with competing bacteria [31].

Using RNA-seq in C. cinerea, we show evidence suggesting a role of the Velvet protein

regulon in sexual development of basidiomycetes. Our data supports the existence of a

conserved transcriptional circuitry in basidiomycetes fruiting bodies consisting of at least 60

orthologous genes probably involved in mushroom development and function. In addition,

our data reveals the existence of two different sets of fungal defense proteins in vegetative

and sexual organs matching the type of competitors and predators by which these structures

are challenged in nature. The transcriptome data is supported by the first partial shotgun

mass spectrometry catalog of proteins present in C. cinerea stage 1 primordia (S1P) and

vegetative mycelium (VM).

Methods

Strains and culture conditions

The dikaryotic, self-compatible C. cinerea strain A43mutB43mut (AB) [32] was grown on 30

mL YMG plates (0.4% yeast extract, 1% malt extract, 50 mM glucose and 1.5% agar) at

37°C in the dark for 96 h and transferred to 25°C, 90% humidity and 12 h photoperiod for

fruiting body production. 1 to 2 mm AB S1P [6] were harvested after 72 h, flash-frozen in

liquid nitrogen and stored at -80°C for later use. C. cinerea AB VM was grown in duplicate

on YMG plates covered with cellophane discs for 96 h at 37°C in the dark and harvested

independently before being flash frozen and stored at -80°C.

Chapter 2 Developmental transcriptomics of C. cinerea

Total RNA extraction

Primordia and mycelia were lyophilized and S1Ps were separated in two pools of 20 mg each. From each VM replicate or S1P pool, 20 mg dry material were lysed in three FastPrep

FP120 homogenization steps of 45 s at 4.5, 5.5 and 6.5 m/s in the presence of 250 mg 0.5 mm glass beads, cooling the samples for 5 min on ice between steps. RNA was extracted using 1 mL Qiazol (Qiagen) and 0.2 mL chloroform ReagentPlus (Sigma-Aldrich). The solution was centrifuged at 12000 xg for 15 min at 4°C; thereafter, RNA from the resultant aqueous phase was washed on-column using the RNeasy Lipid Tissue Mini Kit (Qiagen) and eluted in 60 µL RNase-free water. Concentration and integrity of the purified RNA were determined with a Qubit (1.0) fluorometer (Life Technologies) and a Bioanalyzer 2100

(Agilent), respectively. Samples with a 260/280 nm ratio of 1.8–2.1 and a 28S/18S ratio of

1.5–2 were later used in library construction.

SOLiD 4 library construction

Whole transcriptome libraries from two S1P pools and two VM replicates were produced using MicroPolyA Purist Kit (Ambion) and SOLiD Total RNA-Seq kit (Applied Biosystems).

Briefly, approximately 200 ng/sample poly(A)-positive RNA was enriched using MicroPoly(A)

Purist Kit from 15-20 μg total RNA. Quality and concentration of the extracted poly(A)- positive RNA was re-assessed as described above, and poly(A)-positive RNA was digested with RNase III. Ligation of the adaptor mix and reverse transcription was performed following the manufacturer instructions. cDNA libraries were size selected for 150-250 bp fragments, amplified in 15-18 PCR cycles using barcoded adaptor primers and purified with PureLink

PCR micro kit (Invitrogen). These barcoded cDNA libraries were then amplified by emulsion

PCR from 0.5 pM template. Sequencing beads from the barcoded libraries were pooled and loaded on a SOLiD 4 slide (Applied Biosystems), according to manufacturer’s instructions.

SOLiD ToP Sequencing chemistry was used to produce pair end (50bp+35bp) sequencing reads.

Chapter 2 Developmental transcriptomics of C. cinerea qRT-PCR validation

RNA-seq results were validated by qRT-PCR. Single-stranded cDNA from one biological replicate per sample was synthesized using Transcriptor Universal cDNA Master (Roche) from 2 µg total RNA. 20 µL qRT-PCR reactions were mixed in three technical replicates per primer set and sample, containing 900 nM forward and reverse primers designed to span exon-exon junctions (Table 1), 10 µL 2X FastStart Universal SYBR Green Master (Rox,

Roche) and 1 ng/µL cDNA template. qRT-PCR was performed in a Rotor-Gene 3000

(Corbett Life Science) with the following thermal profile: a hold step at 95°C for 15 min followed by 40 cycles of 95°C for 15 s, 58°C for 30 s and 72°C for 30 s. In order to control the specificity of amplification, the reaction was concluded with a melting curve analysis ramping from 55°C to 99°C in steps of 1°C every 5 s. PCR efficiencies and cycle thresholds were obtained using LinRegPCR 12 [33] and differential expression ratios were calculated by the CT difference formula [34]. Tubulin beta chain (CC1G_04743) was used as a house keeping normalizer. In addition, water or 1 ng/µL RNA were included as negative control reactions. To further validate the significance of the RNA-seq-derived differential expression analysis, the constitutive expression of an array of housekeeping loci commonly used in qRT-PCR normalization [35-41] was verified in the sequencing datasets after library size normalization.

Table 1. qRT-PCR primers used during RNA-seq validation.

Locus Functional annotation Primer name Sequence (5'→3') Tm°C CC1G_04743 Tubulin beta chain TubM1Fw GTCATGTCCGGTATCACCAC 62 CC1G_04743 Tubulin beta chain TubM1Rv GGGAAAGGAACCATGTGGA 61 CC1G_05299 Ricin B-fold protein CC1G_05299FqRTPCR CGGCGACAACCAGCTTTGGGACTTTG 72 CC1G_05299 Ricin B-fold protein CC1G_05299RqRTPCR GACACCGATCCTCCAGACGGATC 68 CC1G_09480 Cospin PICFwqRTPCR ACGTCTTCACCGTCGTGAATGC 67 CC1G_09480 Cospin PICRvqRTPCR TCGACTTGGGTGAAGGTAAAGAGC 66 CC1G_10318 Pore-forming protein CaerofqRTPCR TTTGAGCTCGGGCAGAGGTTAACG 68 CC1G_10318 Pore-forming protein CaeroRqRTPCR AAGCGTGGCCCTCTCCGTGTAAG 70 CC1G_11805 Pore-forming protein CC11805RTPCRFw GGCATCGAACTCGGACAAACGTTCACTTTC 71 CC1G_11805 Pore-forming protein CC11805RTPCRRv AGTTGGTGATCCTCTCCGTGTAAG 65

Chapter 2 Developmental transcriptomics of C. cinerea

Bioinformatic analysis

Fastqc files were used to filter and trim the reads to be mapped. Strand-specific reads were

mapped to the third annotation (September 2010) of the C. cinerea okayama7#130 genome

hosted at the Broad Institute (C. cinerea Sequencing Project, Broad Institute of Harvard and

MIT (http://www.broadinstitute.org/)). SOLiD mapped reads were counted using Cufflinks

[42]. All the sequences were deposited in the ArrayExpress database

(http://www.ebi.ac.uk/arrayexpress/) under the accession number [E-MTAB-1968]. To

determine the percentage of loci showing baseline expression, 5 reads/locus were taken as

a minimal threshold [43]. Library size normalization (scaling method), fold change calculation per locus and fisher exact test comparing mycelia and primordia libraries were performed using the edgeR package [44] using sense-read counts. Fisher exact test p-value ≤ 0.05 and fold change ≥ 8 were the criteria established to classify a locus as differentially expressed in one of the two compared samples. Functional annotation of loci found to be developmentally regulated was explored by PSI-BLAST [45]. Enrichment of annotation terms in differentially

expressed loci was visualized in tag clouds constructed by Wordle (© IBM Corporation) [46];

common tags such as "protein", "hypothetical", "domain", "containing", "family", "putative",

"similar", "probable", "related" and "subunit" were excluded. A complementary functional

annotation clustering was performed with the Database for Annotation, Visualization and

Integrated Discovery (DAVID) 6.7 [47] using default annotation categories and the C. cinerea

okayama7#130 translated genome as background. SignalP 4.1 [48] and TMHMM v. 2.0 [49]

were used to predict the presence of secretion signal and transmembrane helices in

developmentally regulated loci. Presence of nuclear localization signals (NLSs) in members

of the Velvet protein family was assessed with NLStradamus [50]. To analyze the similarity

in the gene expression programs associated to early stages in fruiting body development

across different basidiomycetous fungi, orthologous genes to differentially expressed loci in

C. cinerea S1P were identified by PSI-BLAST (best hit showing an E-value ≤ 0.005) in the

genomes of Schizophyllum commune 4-39/4-40 and Laccaria bicolor S238N-H82.

Expression data were retrieved from the original studies where the gene expression of

Chapter 2 Developmental transcriptomics of C. cinerea vegetative mycelium and S1P or YFB from S. commune [51] and L. bicolor [52], respectively, was measured. First, log2 expression ratios were calculated for stage 1 primordia against monokaryon mycelium from S. commune using tags per million per locus. Similarly, log2 ratios were calculated for "young fruiting bodies" compared to "free living mycelium" (the latter corresponding to the average of two replicates) of L. bicolor from quantile-normalized

Robust multichip average values. A centroid-linkage hierarchical clustering analysis of the computed log2 ratios was performed with Cluster 3.0 and heat maps were generated and visualized using TreeView 3 [53].

A more detailed orthology and expression analysis was carried out for loci known to encode proteins interacting with Velvet domain-containing proteins involved in sexual development and secondary metabolite production in Aspergillus [54]. The sequences of RosA, NosA,

StuA, NsdD, PpoA, VelB, LaeA, KapA FphA and CryA from A. nidulans were used to retrieve the corresponding orthologs from the protein-coding genomes of C. cinerea Okayama 7, S. commune 4-39/4-40 and L. bicolor S238N-H82. Orthologs were defined as the best hits in a

PSI-BLAST showing E-values ≤ 0.005. Progressive multiple sequence alignments for the orthologs present in A. clavatus, A. fumigatus, A. oryzae and A. nidulans, as well as in the three basidiomycetous species considered, were constructed in CLUSTAL W [55]. Finally,

Velvet domain-containing proteins in C. cinerea, S. commune and L. bicolor were identified in Pfam and SMART diagrams were created [56].

LC-MS/MS based shotgun proteomics

Twenty mg lyophilized VM from a single plate or a S1P pool were lysed in a FastPrep FP120 during 40 s at 6 m/s using 200 mg 0.5 mm glass beads and 600 µL PBS supplemented with

1 mM phenylmethanesulfonylfluoride (PMSF) and 1x Complete Protease Inhibitor Cocktail

(Roche). The lysates were centrifuged for 15 min at 16000 xg and 4°C, and the supernatants

(soluble protein extracts) recovered. Soluble protein extracts from VM and S1P were further processed by filter-aided sample preparation (FASP) method as previously described [57]. In

Chapter 2 Developmental transcriptomics of C. cinerea brief, protein extracts were loaded onto an Amicon column equipped with a 10 kDa MWCO membrane (Millipore), reduced with 55 mM dithiothreitol at 37°C for 1 h and alkylated with 65 mM iodoacetamide in the dark at 37°C for 1 h. Reduced and alkylated extracts were digested with sequencing grade porcine trypsin (Roche) for 18 h at 37°C in 25 mM ammonium bicarbonate, pH 8.5. Digested peptide mixtures were collected by centrifugation and dried in a Savant SpeedVac (Thermo Scientific). All samples were de-salted by C18

ZipTip before mass spectrometry analysis.

Samples were analyzed on a LTQ-Orbitrap Velos mass spectrometer (Thermo Fischer

Scientific) coupled to an Eksigent-Nano-HPLC system (Eksigent Technologies). Peptides were suspended in 2.5% acetonitrile and 0.1% formic acid, loaded on a self-made tip column

(75 µm × 80 mm) packed with reverse phase C18 material (AQ,μm 3 200 Å, Bischoff

GmbH) and eluted with 250 nL/min flow rate in a gradient from 3% to 50% of B in 90 min,

97% B in 10 min. One scan cycle comprised a full scan MS survey spectrum, followed by up to 20 sequential CID MS/MS on the most intense signals above a threshold of 1500. Full- scan MS spectra (300–2000 m/z) were acquired in the FT-Orbitrap at a resolution of 60000 at 400 m/z, while CID MS/MS spectra were recorded in the linear ion trap. CID was performed with a target value of 1E4 in the linear trap, collision energy at 35 V, Q value at

0.25 and activation time at 30 ms. AGC target values were 5E5 for full FTMS scans and 1E4 for ion trap MSn scans. For all experiments, dynamic exclusion was used with one repeat count, 15 s repeat duration, and 60 s exclusion duration. For quantitation, each sample was measured in technical triplicates using the same parameters.

All MS data from VM and S1P were converted to a peak list and searched against the C. cinerea okayama7#130 proteome (predicted transcript translation) hosted at the Broad

Institute (C. cinerea Sequencing Project, Broad Institute of Harvard and MIT

(http://www.broadinstitute.org/)) using the Mascot search engine (version 2.3) considering variable modifications: carbamidomethylation on cysteine and oxidation on methionine. The

Chapter 2 Developmental transcriptomics of C. cinerea tolerance of mass accuracy of MS and MS/MS was 8 ppm and 0.6 Da. The false discovery rate of proteome dataset was 1% and the score of each protein exceeded 30.

Cloning and recombinant expression of CC1G_11805-encoding cDNA

The gene encoding CC1G_11805 was amplified from C. cinerea S1P-derived cDNA using the forward and reverse primers 5'-CCAGCTTAAAGGAGTCACAAGG-3' and 5'-

AACGTTCAACGCCCAGCCAC-3', respectively, with a Pfu DNA polymerase. 3' adenines were added to the PCR fragment before ligation to the DNA amplification vector pGEM-T

Easy (Promega). This pGEM-CC1G_11805 construct was used as a template to add NdeI and BamHI restriction sites using the primers 5'-

GGGGGGCATATGTCTCAAGCAGGGATCACAC-3' and 5'-

GGGGGGGGATCCTCAGATACGCCCGATGACTTC-3', respectively. The resulting product was digested with NdeI and BamHI in 2x Tango buffer (Thermo Scientific), ligated to a pre- digested pET24 expression vector (EMD Millipore), and used to transform chemocompetent

E. coli BL21. Transformed colonies were selected on LB plates containing 50 µg/mL

Kanamycin. CC1G_11805 recombinant expression and solubility were assessed as follows: pET24-CC1G_11805-containing E.coli BL21 was grown in LB broth supplemented with 50

µg/mL Kanamycin up to OD600: 0.5; thereafter, the cultures were divided in two separated flasks and protein expression was induced for 18 h at 24°C by adding 1 mM Isopropyl β-D-1- thiogalactopyranoside (IPTG, final concentration) to only one of the two subcultures. Cells were harvested, suspended in 1 mL PBS containing 1 mM phenylmethanesulfonylfluoride

(PMSF) and lysed in a single FastPrep FP120 homogenization step of 35 s at 6 m/s in the presence of 1 g 0.1 mm glass beads. The whole cell protein extract (WCE) was centrifuged for 5 min at 5000 xg and 4°C (low speed centrifugation, LS) to remove cell debris and the supernatant transferred to a fresh tube to be centrifuged again for 30 min at 14000 xg and

4°C (high speed centrifugation, HS). Samples were collected from the whole cell extracts

(WCE and WCE+IPTG) and the two centrifugation supernatants (LS and HS), mixed with

Laemmli buffer and denatured at 95°C for 5 min to be loaded on a 12% SDS-PAGE.

Chapter 2 Developmental transcriptomics of C. cinerea

Caenorhabditis elegans toxicity assay

In order to evaluate the developmental toxicity of CC1G_11805 to nematodes, a C. elegans

toxicity assay was performed as previously described [58]. In brief, C. elegans N2 and C.

elegans pmk-1 (km25) (kindly provided by M. O. Hengartner) were grown on NGM plates (50

mM NaCl, 2.5 g/L bacteriological peptone, 13 mM cholesterol and 1.7% agar) pre-seeded

with E. coli OP50. Eggs were obtained by bleaching gravid hermaphrodites in 15 mL conical

bottom tubes using a solution containing 0.5 N NaOH and 1% NaClO for 10 min and

washing twice in 10 mL distilled deionized water (ddH2O). Clean eggs were transferred to a

1.5% agar plate and hatched for 18 h at 20°C. L1 larvae were collected in PBS, counted and

adjusted to 1.5 larvae/µL. Approximately 30 L1 larvae/replicate (4 replicates/treatment) were

mixed in 200 µL PBS with OD600: 2 IPTG-pre-induced E. coli BL21 expressing CC1G_11805

in flat bottom 96 well-plates. As a negative control, IPTG-pre-induced E. coli BL21 cells

transformed with empty vector pET24 were used. Worms were incubated for 48 h at 20°C

and the number of individuals reaching each developmental stage or dying was counted. A

Mann Whitney test between the empty vector pET24 control and CC1G_11805 was run to

test the statistical significance of the results observed.

Aedes aegypti toxicity assay

CC1G_11805 toxicity towards A. aegypti Rockefeller (kindly provided by W. Rudin and P.

Müller) was tested as previously described [58]. In brief, 600-800 eggs were hatched in a

glass petri dish containing 200 mL deionized water and 30 mg ground fish food for 20 h at

28°C in the dark. Larvae were transferred to fresh 800 mL deionized water supplemented

with 50 mg ground fish food and incubated for 10 h at 28°C in order to obtain synchronized

L2 larvae. Ten L2 larvae/replicate (4 replicates/treatment) were starved in 100 mL fresh

deionized water in 100 mL Schott flasks for 6 h at 28 °C before adding 1 mL OD600: 20 (Final

OD600: 0.2) IPTG-pre-induced E. coli BL21 (empty vector pET24, pET24-CGL2 [18] or

pET24-CC1G_11805). Larvae were incubated for 96 h at 28°C in the dark and the surviving

individuals were counted. Percentages of surviving larvae/treatment were calculated. A

Chapter 2 Developmental transcriptomics of C. cinerea

Dunn's multiple comparison test between the empty vector pET24 control and the different

treatments was run to test the statistical significance of the results observed.

Results

Differential gene expression during fruiting body development in C. cinerea.

Four different cDNA libraries were sequenced with a final 7.32 Gb mapped data output.

Approximately 95% of the open reading frames (ORFs) in the genome of C. cinerea

A43mutB43mut (AB) were transcribed using five reads/ORF as the minimal threshold for a

locus to be considered as expressed (Table 2).

Table 2. General features of the C. cinerea S1P and VM transcriptomes.

Library size (reads) Biological replicate 1 Biological replicate 2 Expression (%)* S1P 26243754 25653679 95.55 VM 29374279 23269631 95.13 Average SOLiD read size (b) 70 Sequenced material (Gb) 7.32 * Mean percentage of annotated ORFs detected to be expressed using a minimal threshold

5 reads/ORF.

Differential gene expression at the RNA level between S1P and VM was examined. Eleven

percent of the annotated ORFs in the C. cinerea genome were found to be differentially

expressed, 795 (6%) and 679 loci (5%) in VM and S1P, respectively, using fold change 8 and Fisher's exact test p-value ≤ 0.05 as thresholds (Figure 1). The number of differentially transcribed loci in these two developmental stages increased to 2522 in VM and 3209 in

S1P when a fold change threshold of 2 was set, corresponding to approximately 45%

annotated ORFs in the genome of C. cinerea.

Chapter 2 Developmental transcriptomics of C. cinerea

Figure 1. Differential gene expression during early fruiting body development in C.

cinerea. (A) Volcano plot illustrating the relative expression (log2(S1P/VM)) and statistical

significance of 13342 loci of the genome of C. cinerea between VM and S1P. Fisher's exact test -log10 p-value ≥ 1.3 (p-value ≤ 0.05) and log2(S1P/VM) +/- 3 (8 fold change) were used as thresholds of differential gene expression. Grey boxes comprise genes significantly up- regulated in S1P (right) or VM (left). Tag clouds showing enriched PSI-BLAST functional annotation terms from up-regulated loci in S1P (B) and VM (C) were produced in Wordle (©

IBM Corporation) after removing frequently appearing tags.

To validate these data, qRT-PCR of four selected loci (CC1G_10318, CC1G_09480,

CC1G_05299 and CC1G_11805) was performed. Although S1P/VM expression ratios did

not exactly match those observed by RNA-seq, qRT-PCR results showed the same trend of

differential gene expression for these loci during C. cinerea development as observed by

SOLiD RNA-seq (Figure 2). In addition, we verified that the expression values of reported

housekeeping loci commonly used in qRT-PCR normalization were in a range indicative of

constitutive expression (-2 ≤ log2(S1P/VM) ≤ 2) (Table 3).

Chapter 2 Developmental transcriptomics of C. cinerea

Figure 2. RNA-seq data validation by qRT-PCR. The expression of four selected genes was validated by qRT-PCR. A comparable relative expression pattern was found for all the genes evaluated in S1P and VM with both techniques. RNA-seq data corresponds to the mean log2(S1P/VM) of two biological replicates. qRT-PCR data show the mean log2(S1P/VM) of three technical replicates from a single biological replicate of S1P and VM.

Bars correspond to standard deviations. Dashed lines indicate the differential gene expression thresholds selected for this study (log2(S1P/VM) = +/-3)

Chapter 2 Developmental transcriptomics of C. cinerea

Table 3. Reference housekeeping loci are constitutively expressed in C. cinerea

A43mutB43mut.

Reads/kb Locus Functional annotation VM1 VM2 S1P1 S1P2 log2(S1P/VM) Reference CC1G_06184 Beta tubulin 3187 4296 2746 3543 -0.2510 Wan H 2011 CC1G_04743 Tubulin beta chain 57352 69507 140002 168645 1.2827 Wan H 2011 CC1G_07639 Histone H2B 49428 50071 144222 173258 1.6739 Ferreira 2012 CC1G_03523 Histone H2B 44912 38795 77045 79233 0.9007 Ferreira 2012 CC1G_13048 Actin 677 624 792 878 0.3605 Huggett 2005 CC1G_08232 Actin 96495 116431 112483 108304 0.0523 Huggett 2005 CC1G_09116 GADPH 119239 151363 40820 40636 -1.7321 Huggett 2005 CC1G_09117 GADPH 205 187 119 121 -0.7087 Huggett 2005 CC1G_11833 Ubiquitin 99610 150276 185548 200736 0.6284 Silveira 2009 CC1G_00876 Ubiquitin 80981 91185 127562 128692 0.5738 Silveira 2009 CC1G_03676 Ubiquitin C 353488 272284 149754 176153 -0.9412 Silveira 2009 CC1G_09572 Cyclophilin 333724 296676 103503 92140 -1.6880 Langnaese 2008 CC1G_15352 Ribosomal protein S27a 255 232 141 150 -0.7416 de Oliveira 2011 CC1G_04355 Ribosomal protein L19 80930 74132 55940 58059 -0.4438 de Oliveira 2011 CC1G_00758 Ribosomal protein L11 58743 49275 64729 61966 0.2301 de Oliveira 2011 CC1G_13649 Ribosomal protein L32 83383 81823 75132 78869 -0.1013 de Oliveira 2011 CC1G_03927 Hsp90 16541 22271 22405 31048 0.4618 Aursnes 2011

Enrichment of PSI-BLAST-derived functional annotation terms was visualized using Wordle

(426 loci); nonetheless, there was a significant enrichment of N-methyltransferase (histone- lysine N-methyltransferase) tags in S1P (7 loci). Enrichment of functional annotation terms also shows that different sets of cytochromes, kinases, dehydrogenases, transporters and hydrophobins are specifically expressed in vegetative mycelium or young fruiting bodies

(Figure 1). Functional annotation clustering using DAVID Bioinformatics Resources 6.7 [47] confirmed the enrichment of protein methyltransferases and hydrophobins in S1P with 11 and 6 fold enrichment compared to the occurrence of these functional categories in the genome of C. cinerea.

Tissue-specific expression of C. cinerea defense proteins

Several cytoplasmic defense lectins and protease inhibitors such as C. cinerea lectin 2

(CCL2, CC1G_11781) [59], Coprinopsis galectin 1 (CGL1, CC1G_05003) [60] and two paralogous serine protease inhibitors from Coprinopsis (Cospin; CC1G_09479 and

CC1G_09480) [20] were found to be highly up-regulated in C. cinerea S1P (Table 4).

Chapter 2 Developmental transcriptomics of C. cinerea

Table 4. S1P-specific defense loci.

Locus Fold S1P/VM Functional annotation *p-value †SignalP ‡TMHMM CC1G_09480 2426 Cospin1 1.26E-305 N N CC1G_11781 1939 CCL2 4.64E-302 N N CC1G_09479 692 Cospin2 2.38E-132 N N CC1G_12219 237 Related to Velvet A protein 6.42E-43 N N CC1G_07937 83 Ricin B-fold protein 1.08E-148 Y N CC1G_11778 69 CCL1 7.80E-137 N N CC1G_10318 68 Pore-forming protein 8.08E-91 N N CC1G_07956 66 Peptidoglycan-binding domain 1 protein 1.70E-48 Y 1 CC1G_14321 35 Hemolysin 9.81E-77 N 3 CC1G_08484 34 Cercosporin toxin biosynthesis protein 4.71E-115 N N CC1G_06959 31 Thaumatin-like protein 3.50E-79 Y N CC1G_13099 15 Peptidoglycan-binding domain 1 protein 4.40E-17 N N CC1G_05003 12 CGL1 7.13E-37 N N CC1G_11123 11 Toxin-antitoxin system, toxin component 1.84E-33 N N CC1G_11246 11 Ricin B-fold protein 7.70E-28 Y N CC1G_11805 11 Pore-forming protein 1.69E-40 N N * Fisher's exact test.

† N or Y indicates the lack or presence of a signal peptide.

‡ N indicates a lack of transmembrane helices, while a number corresponds to the amount

of transmembrane helices predicted in the ORF.

Out of these previously characterized genes coding for proteins with nematotoxic and

insecticidal activity, Cospin and CCL2 were found to be among the top 50 most highly transcribed and differentially expressed loci in S1P. Among these, two loci, CC1G_10318 and CC1G_11805, encoding homologous proteins with a predicted aerolysin/ETX pore- forming domain as well as loci encoding proteins with suspected antibacterial and antifungal function including two peptidoglycan binding proteins, the toxin component of a bacterial toxin-antitoxin system and a Thaumatin-like protein were found. In contrast to S1P-specific defense proteins, defense proteins specifically up-regulated in VM were mainly secreted.

These proteins included three putative lysozymes, several proteins containing a CFEM

domain (PF05730) whose structure resembles cysteine-stabilized antibacterial peptides, and

two representatives of the cerato-platanin family of secreted proteins (Table 5).

Chapter 2 Developmental transcriptomics of C. cinerea

Table 5. VM-specific defense loci.

Locus Fold VM/S1P PSI Blast *p-value †SignalP ‡TMHMM CC1G_05299 2076 Ricin B-fold protein 2.95E-240 N N CC1G_10614 1481 CFEM domain-containing protein 9.18E-166 Y N CC1G_15645 851 CFEM domain-containing protein 1.51E-194 Y N CC1G_09154 639 Cerato-platanin protein 6.24E-233 Y N CC1G_13813 527 CFEM domain-containing protein 6.90E-195 Y N CC1G_05638 67 Peptidoglycan-binding domain 1 protein 1.19E-119 Y 1 CC1G_05246 42 Ricin B-fold protein 7.71E-125 Y N CC1G_09155 24 Cerato-platanin protein 3.27E-75 Y N CC1G_09421 21 Terpenoid synthase 5.98E-27 N N CC1G_11847 17 Lysozyme 2.00E-08 Y N CC1G_08066 16 Ricin B-fold protein 1.01E-45 Y N CC1G_08310 13 Lysozyme 6.90E-09 Y N CC1G_15739 10 Ricin B-fold protein 1.54E-59 Y N CC1G_03046 8 Lysozyme 3.68E-05 Y N * Fisher's exact test.

† N or Y indicates the lack or presence of a signal peptide.

‡ N indicates a lack of transmembrane helices, while a number corresponds to the amount

of transmembrane helices predicted in the ORF.

Latter protein family is expanded in basidiomycetes and has recently been implicated in

interactions of dikaryotic fungi with other organisms [61, 62]. During sexual development on

herbivore dung, C. cinerea is exposed to a succession of antagonists (predators and competitors) colonizing this substrate. The differential expression of cytoplasmic and secreted defense proteins in S1P and VM, respectively, might reflect the prevalent types of antagonists with which these tissues of C. cinerea are confronted.

The S1P-specific aerolysin/ETX pore-forming domain-containing protein CC1G_11805

is toxic to nematode and insect larvae

CC1G_11805, CC1G_10318 and CC1G_08369 encode three homologous 30-40 kDa

proteins containing a predicted aerolysin/ETX pore-forming domain (Figure 3A). This

domain is homologous to the one present in the insecticidal epsilon toxin (ETX) from

Clostridium perfringens [63] and distantly related to aerolysin pore-forming toxins [64]. In

order to test the significance of the up-regulation of CC1G_11805 and CC1G_10318 in S1P

with regard to fungal defense, we cloned and recombinantly expressed CC1G_11805 in

Chapter 2 Developmental transcriptomics of C. cinerea

Escherichia coli and assessed the toxicity of the protein by feeding the recombinant bacteria to nematode and insect larvae as described previously [58]. Results in Figure 3B show that

CC1G_11805 was expressed in soluble form in E. coli. Feeding of CC1G_11805-expressing bacteria to L2 larvae of the mosquito Aedes aegypti lead to their death after 96 h (Figure

3C). Vector control-containing and fungal lectin-expressing E. coli were used as negative and positive controls, respectively, in these experiment [18, 29]. Similarly, feeding of

CC1G_11805-expressing bacteria was found to significantly impair larval development of the

C. elegans N2 wildtype and pmk-1(km25) mutant strains (Figure 3D); in agreement with the previously reported higher susceptibility of latter strain to different kinds of abiotic and biotic stresses including other nematotoxic fungal defense proteins [60], CC1G_11805 was shown to be more toxic to C. elegans pmk-1(km25) than to N2. These results support previous observations [18] showing that the expression of defense proteins directed against nematodes and insects is significantly increased in C. cinerea sexual organs.

Chapter 2 Developmental transcriptomics of C. cinerea

Figure 3. The S1P-specific aerolysin/ETX pore-forming domain-containing protein

CC1G_11805 is toxic for C. elegans and A. aegypti. (A) Multiple sequence alignment of

the aerolysin/ETX pore-forming domain-containing proteins C. perfringens epsilon toxin

(Cper_etx), CC1G_11805, CC1G_10308 and CC1G_08369. (B) Expression of soluble

CC1G_11805 in E. coli BL21. Whole cell protein extracts in PBS were produced from non-

induced (WCE) and induced (WCE+IPTG) E. coli BL21/pET24-CC1G_11805 cultures.

Supernatants from consecutive low (LS) and high (HS) speed centrifugations were collected

and run on a 12% SDS-PAGE to evaluate CC1G_11805 expression and solubility.

CC1G_11805-mediated toxicity for A. aegypti (C) and C. elegans (D) larvae was assessed

as described previously [58]. E. coli BL21 expressing the previously characterized fungal

Chapter 2 Developmental transcriptomics of C. cinerea

lectin CGL2 was used as positive control for toxicity against A. aegypti Rockefeller. IPTG-

induced E. coli bearing an empty pET24 vector (EV) was used as a negative control.

Columns represent the mean percentage of surviving insect larvae (C), or worms either

dying or reaching the indicated developmental stage (D) from 4 replicates each. SDs are

shown as error bars. Dunn's multiple comparisons were used to test the statistical

significance of the toxicity observed in the A. aegypti assay. Mann Whitney test was used to

compare the percentage of worms reaching each developmental stage when treated with EV

or CC1G_11805. *: 0.01 < p-value < 0.05; **: 0.001 < p-value < 0.01.

Differential expression of loci involved in sexual development of filamentous fungi

Various genes were recently described as playing a role at different stages of fruiting body

formation of C. cinerea [10-17]. With the exception of the cyclopropane fatty acid synthase

cfs1, none of these loci were found to be differentially transcribed during C. cinerea sexual

development (Table 6, Figure 4).

Table 6. Most genes necessary for fruiting body formation in C. cinerea are not up-regulated

in S1P.

Locus Fold S1P/VM Gene *p-value Mutant phenotype Reference CC1G_03287 0.8 rmt1 1.43E-01 No clamp connections Nakazawa 2010 CC1G_00975 1.4 ubc2 1.75E-02 No hyphal knot formation Nakazawa 2011 CC1G_15539 1.1 snf5 3.98E-01 Blocked at hyphal knot Ando 2013 CC1G_11387 64.0 cfs1 5.25E-142 Blocked at initials Liu 2006 CC1G_06825 1.2 dst2 9.66E-02 Blocked at stage 2 primordia Kamada 2010 CC1G_08609 2.6 dst1 1.21E-13 Blocked at stage 2 primordia Kamada 2010 CC1G_10193 1.5 eln3 1.40E-03 Blocked at immature mushroom Arima 2004 CC1G_04713 2.3 eln3 5.18E-08 Blocked at immature mushroom Arima 2004 CC1G_06451 2.5 eln3 9.03E-13 Blocked at immature mushroom Arima 2004 CC1G_01334 0.8 exp1 3.65E-02 Blocked before mushroom decay Muraguchi 2008 * Fisher's exact test.

The Velvet protein regulon plays a major role in the control of sexual vs. asexual development in the ascomycete Aspergillus nidulans [54, 65]; therefore, we took a closer look at the expression pattern of C. cinerea orthologs of the Velvet protein family and the proteins regulated by it [54, 65]. CC1G_12219, a Velvet domain-containing protein bearing an NLS is highly induced in S1P (S1P/VM = 237). In total, three out of six Velvet domain-

Chapter 2 Developmental transcriptomics of C. cinerea

containing proteins encoded in the C. cinerea genome (CC1G_12219, CC1G_06962 and

CC1G_14883) were found to be developmentally regulated (Figure 5). Among the other

members of the Velvet protein regulon, two homologs of the VeA-regulated activator NsdD

(CC1G_06391 and CC1G_06265) were also up-regulated in S1P (S1P/VM = 14 and 2.4,

respectively). Moreover, the C. cinerea homeodomain transcription factor STE-12

(CC1G_02207) showed a moderate up-regulation in primordia (S1P/VM = 2). Finally,

CC1G_07060, an ortholog of the A. nidulans repressor of sexual development rosA, was 34 fold up-regulated in VM compared to S1P. Taken together, the differential expression of several members of the Velvet protein regulon suggests that these genes may play a role in sexual development of basidiomycetes.

Figure 4. Fruiting body development in C. cinerea. Gene mutations preventing fruiting body development at different stages in C. cinerea are shown in blue. White and black sections indicate light and dark periods, respectively, corresponding to 12 h each. Numbers indicate the measured fold (S1P/VM). A mutation in the S1P-specific locus cfs1 stops sexual development at the initials stage. Figure adapted from [6].

Chapter 2 Developmental transcriptomics of C. cinerea

Figure 5. Differential transcription comparison between genes encoding Velvet domain-containing proteins from three different basidiomycetes. Amino acid

sequences corresponding to the full set of Velvet domain-containing proteins encoded in the

genomes of C. cinerea, L. bicolor and S. commune were retrieved from Pfam (PF11754).

SMART domain architecture diagrams are shown in (A). (B) Loci expression in S1P (C. cinerea and S. commune) or YFB (L. bicolor) relative to VM for all the velvet domain- containing proteins in the three basidiomycetes compared. N/E: No expression detected.

Chapter 2 Developmental transcriptomics of C. cinerea

Conserved transcriptional circuitry during fruiting body development among basidiomycetous fungi

In order to explore a hypothetical conserved gene expression circuitry during the formation of SR structures in basidiomycetes, a comparative transcriptomic analysis by hierarchical clustering including C. cinerea, S. commune [51] and L. bicolor [52] was performed.

Orthologous genes in S. commune and L. bicolor were assigned from a PSI-BLAST search

(hits with the lowest E-value ≤ 0.005) to 446 and 378 up- and down-regulated loci in C. cinerea S1P, respectively. Massively parallel signature sequencing tags/million from S. commune monokaryotic mycelium and stage 1 primordia (ScS1P); as well as L. bicolor free- living monokaryotic mycelium (2 replicates) and young fruiting bodies (LbYFB, one replicate) robust multichip average values were retrieved from original studies [51, 52]. log2 expression ratios were calculated using monokaryotic mycelium expression values from these conserved loci as denominators and a centroid linkage hierarchical clustering analysis was computed (Figure 6). Four clusters corresponding to 37 up-regulated loci in early sexual development in basidiomycetes were identified (Pearson correlation coefficients:

0.95, 0.99, 0.99 and 0.99, respectively). In addition, two different gene groups (29 loci) were found to be down-regulated in early stages of fruiting body formation when compared to vegetative mycelium of the three species (Pearson correlation coefficients: 0.99 and 0.94, respectively) (Table 7).

Chapter 2 Developmental transcriptomics of C. cinerea

Figure 6. Conserved expression pattern of some gene clusters during fruiting body formation among three different basidiomycetous species. Comparative transcriptome

analysis of early sexual development in S. commune 4-39/4-40, C. cinerea AB and L. bicolor

S238N-H82 was performed. Orthologous genes (best PSI-BLAST hits with E-values ≤ 0.005) to only 446 and 378 up- and down-regulated loci in S1P, respectively, were found in S. commune 4-39/4-40 and L. bicolor S238N-H82. (A) Fruiting body transcriptome data were retrieved in the form of log2 ratios for these loci from the raw data sets and analyzed by centroid-linkage hierarchical clustering. The red sidelines mark the presence of 6 loci clusters which are consistently up (1-4)- or down (5 and 6)-regulated in young fruiting bodies (YFB) and stage 1 primordia (S1P) of the three species compared. (B) Close-up view of the up- and down-regulated clusters highlighted by red bars in (A). Orthologous genes in

C. cinerea, S. commune and L. bicolor corresponding to the numbered clusters, as well as

the functional annotation of these loci and the cluster-associated Pearson correlation

coefficients can be found in Table 7.

Chapter 2 Developmental transcriptomics of C. cinerea

Table 7. Up- and down-regulation of orthologous loci is conserved in stage 1 primordia and

young fruiting bodies of three basidiomycete species.

Cluster C. cinerea locus S. commune locus L. bicolor locus Functional annotation* 1(PC†: 0.95) CC1G_01874 SCHCODRAFT_73392 LACBIDRAFT_301146 hypothetical protein CC1G_02466 SCHCODRAFT_109665 LACBIDRAFT_297494 hypothetical protein CC1G_00973 SCHCODRAFT_83399 LACBIDRAFT_293318 chitin deacetylase CC1G_10475 SCHCODRAFT_57566 LACBIDRAFT_295571 aromatic peroxygenase precursor CC1G_14095 SCHCODRAFT_48101 LACBIDRAFT_294461 hypothetical protein CC1G_03094 SCHCODRAFT_52606 LACBIDRAFT_315775 hypothetical protein CC1G_06074 SCHCODRAFT_76530 LACBIDRAFT_293318 carbohydrate esterase family 4 protein CC1G_10044 SCHCODRAFT_108884 LACBIDRAFT_306386 related to S.pombe pac2 protein CC1G_00753 SCHCODRAFT_234371 LACBIDRAFT_305130 hypothetical protein CC1G_14786 SCHCODRAFT_52306 LACBIDRAFT_246776 MFS nicotinic acid transporter Tna1 CC1G_01301 SCHCODRAFT_72462 LACBIDRAFT_309640 hypothetical protein CC1G_05059 SCHCODRAFT_52448 LACBIDRAFT_308735 symbiosis-related protein CC1G_11437 SCHCODRAFT_67374 LACBIDRAFT_317173 putative aquaporin 6 CC1G_08178 SCHCODRAFT_237423 LACBIDRAFT_323328 thioesterase family protein CC1G_08180 SCHCODRAFT_237423 LACBIDRAFT_323328 thioesterase CC1G_01879 SCHCODRAFT_257636 LACBIDRAFT_326397 mei2 protein Piriformospora CC1G_05223 SCHCODRAFT_70203 LACBIDRAFT_250946 DUF1275 domain protein CC1G_12219 SCHCODRAFT_28806 LACBIDRAFT_317102 related to velvet A protein CC1G_10750 SCHCODRAFT_86141 LACBIDRAFT_184665 glutathione S-transferase CC1G_04060 SCHCODRAFT_13677 LACBIDRAFT_180892 hydrophobin-251 CC1G_10471 SCHCODRAFT_232646 LACBIDRAFT_295571 aromatic peroxygenase precursor CC1G_.12330 SCHCODRAFT_16019 LACBIDRAFT_256021 S-layer domain-containing protein 2 (PC: 0.99) CC1G_00069 SCHCODRAFT_48115 LACBIDRAFT_312065 C factor cell-cell signaling protein CC1G_00217 SCHCODRAFT_46102 LACBIDRAFT_323571 Non-Catalytic module family EXPN protein CC1G_03515 SCHCODRAFT_82058 LACBIDRAFT_328112 hydrophobin-like protein CC1G_04659 SCHCODRAFT_110821 LACBIDRAFT_298314 hypothetical protein CC1G_05652 SCHCODRAFT_232455 LACBIDRAFT_309898 TKL/TKL-ccin protein kinase CC1G_09209 SCHCODRAFT_110821 LACBIDRAFT_298284 BTB/POZ domain containing protein CC1G_10966 SCHCODRAFT_258323 LACBIDRAFT_236299 rCop c3 3 (PC: 0.99) CC1G_02571 SCHCODRAFT_76530 LACBIDRAFT_293318 chitin deacetylase CC1G_03037 SCHCODRAFT_53388 LACBIDRAFT_239714 monocarboxylate transporter CC1G_03320 SCHCODRAFT_39718 LACBIDRAFT_145110 Spo11 CC1G_09616 SCHCODRAFT_235946 LACBIDRAFT_310809 prenyl cysteine carboxyl methyltransferase CC1G_06563 SCHCODRAFT_46720 LACBIDRAFT_243581 exo-beta-1,3-glucanase CC1G_12688 SCHCODRAFT_55636 LACBIDRAFT_296585 aldo/keto reductase 4 (PC: 0.99) CC1G_03949 SCHCODRAFT_258034 LACBIDRAFT_298461 salicylate hydroxylase CC1G_05817 SCHCODRAFT_258034 LACBIDRAFT_298461 salicylate 1-monooxygenase 5 (PC: 0.99) CC1G_06868 SCHCODRAFT_53362 LACBIDRAFT_182606 endo-1,3(4)-beta-glucanase CC1G_07550 SCHCODRAFT_56140 LACBIDRAFT_291657 lipase CC1G_05781 SCHCODRAFT_72461 LACBIDRAFT_291413 OrdA protein CC1G_08860 SCHCODRAFT_80181 LACBIDRAFT_172283 cation/H+ exchanger CC1G_09431 SCHCODRAFT_49922 LACBIDRAFT_190903 O-methylsterigmatocystin oxidoreductase CC1G_11000 SCHCODRAFT_107054 LACBIDRAFT_310382 hypothetical protein 6 (PC: 0.94) CC1G_02440 SCHCODRAFT_103563 LACBIDRAFT_308947 ras GEF CC1G_07544 SCHCODRAFT_80832 LACBIDRAFT_243185 DUF89-containing protein CC1G_05410 SCHCODRAFT_104205 LACBIDRAFT_301153 hypothetical protein CC1G_02628 SCHCODRAFT_82883 LACBIDRAFT_327335 C2h2 conidiation transcription factor FlbC CC1G_11894 SCHCODRAFT_68168 LACBIDRAFT_296675 bas1, putative CC1G_05329 SCHCODRAFT_85265 LACBIDRAFT_300118 hypothetical protein CC1G_09061 SCHCODRAFT_27314 LACBIDRAFT_308057 MNNG and nitrosoguanidine resistance protein CC1G_02380 SCHCODRAFT_77933 LACBIDRAFT_314347 Clavaminate synthase-like protein CC1G_05793 SCHCODRAFT_85265 LACBIDRAFT_300118 hypothetical protein CC1G_14014 SCHCODRAFT_49922 LACBIDRAFT_248655 O-methylsterigmatocystin oxidoreductase CC1G_11136 SCHCODRAFT_232299 LACBIDRAFT_294457 hypothetical protein CC1G_00780 SCHCODRAFT_106836 LACBIDRAFT_305061 BGP, partial CC1G_03120 SCHCODRAFT_83385 LACBIDRAFT_307143 endoglucanase II CC1G_02257 SCHCODRAFT_46132 LACBIDRAFT_295167 metalloprotease CC1G_04116 SCHCODRAFT_75425 LACBIDRAFT_179508 glutathione S-transferase Gst3 CC1G_11081 SCHCODRAFT_75425 LACBIDRAFT_179508 glutathione S-transferase CC1G_12752 SCHCODRAFT_50449 LACBIDRAFT_253329 cytochrome P450 CC1G_12253 SCHCODRAFT_269973 LACBIDRAFT_296037 Nrg1-like Zn-finger transcription factor CC1G_11786 SCHCODRAFT_58473 LACBIDRAFT_315853 hypothetical protein CC1G_09938 SCHCODRAFT_80558 LACBIDRAFT_330070 Ferritin/ribonucleotide reductase-like-protein CC1G_02724 SCHCODRAFT_255861 LACBIDRAFT_295183 hypothetical protein CC1G_05802 SCHCODRAFT_65657 LACBIDRAFT_256254 indole-3-acetaldehyde dehydrogenase CC1G_15739 SCHCODRAFT_76887 LACBIDRAFT_146952 Ricin B fold protein *Functional annotation as determined by PSI-BLAST.

†Pearson correlation coefficient

Chapter 2 Developmental transcriptomics of C. cinerea

This hierarchical clustering approach of gene expression in orthologous loci during sexual

development showed that loci involved in core meiotic functions, such as mei2

(CC1G_01879) and spo11 (CC1G_03320), are up-regulated during the early stages of

fruiting body formation in basidiomycetes.

Given the up-regulation of a conserved Velvet-domain containing protein in primordia of the

analyzed basidiomycetes, the conservation and expression pattern of Velvet-interacting

proteins previously characterized in Aspergillus [54] was examined. With the exception of

cryA, orthologs to these Velvet-interacting proteins can be found in C. cinerea, L. bicolor and

S. commune (Additional file 10). Contrary to the conserved transcriptional regulation

observed for genes encoding Velvet domain-containing proteins, little conservation in

expression is evident for the orthologs to rosA/nosA, stuA, nsdD, ppoA, laeA or fphA (Figure

7). Nevertheless, conserved down-regulation during sexual development is shown for velB and kapA, suggesting that a fraction of the Velvet-associated regulon described for ascomycetes might be playing a role during sexual development in basidiomycetes (Figure

7). Similarly, down-regulation of transcription factors such as flbC (a paralog to C2H2 known

to regulate development in A. nidulans [66]), nrg1 and bas1 during early mushroom formation was conserved between the three basidiomycetous species.

Chapter 2 Developmental transcriptomics of C. cinerea

Figure 7. Differential transcription comparison between genes coding for Velvet- associated proteins from three different basidiomycetes. C. cinerea, L. bicolor and S. commune loci homologous to those encoding VelvetA-associated proteins in the ascomycete Aspergillus clavatus (shown on top) were identified by PSI-BLAST (Best hit showing an E-value < 0.005). Differential expression in S1P or YFB relative to VM is shown for the orthologs identified. log2(S1P or YFB/VM) > 0 indicates increased expression in S1P

(C. cinerea or S. commune) or YFB (L. bicolor). On the contrary, a log2(S1P or YFB/VM) < 0 represents a decreased expression in S1P or YFB. Blue and white locus IDs differentiate neighboring groups of orthologs in the chart. Expression of velB and kapA is conserved during sexual development among basidiomycetes. N/E: No expression detected.

Transcription factor mutants (Bri1, Hom1, Gat1, Fst3, C2h2, Fst4 and Hom2) altering normal sexual development in S. commune have been described [67, 68]. Although orthologs of these transcription factors are present in C. cinerea and L. bicolor, the respective genes do not show a conserved expression pattern, indicating that there might be differences in the interplay between these factors during fruiting body formation between different basidiomycetes (Figure 8).

Chapter 2 Developmental transcriptomics of C. cinerea

Figure 8. Differential transcription comparison between genes coding for transcription factors involved in sexual development in S. commune from three different basidiomycetes. Orthologs of transcription factors involved in sexual development in S. commune were identified by PSI-BLAST in the genomes of C. cinerea and L. bicolor. log2((S1P or YFB)/VM) < 0: Down-regulation in S1P or YFB; log2((S1P or YFB)/VM) > 0: up- regulation in S1P or YFB. Domain architecture for each ortholog in the three species compared is shown in the right hand panel.

Chapter 2 Developmental transcriptomics of C. cinerea

Shotgun MS analysis of PBS-soluble VM and S1P proteins

In order to confirm protein expression of some of the genes found to be differentially

expressed by RNA-seq, we assessed the PBS-soluble C. cinerea proteome in VM and S1P by LC-MS. The analysis detected peptides corresponding to a total of 493 proteins in the samples, including 41, 141 and 311 proteins in S1P, VM or both, respectively. This analysis is highly biased towards abundant and soluble proteins and is likely to have failed to detect peptides from most of the loci identified by RNA-seq. Nevertheless, this method allowed us to confirm the translation of 12 transcripts up-regulated in S1P and 50 transcripts up- regulated in VM into proteins in the respective tissues.

Discussion

Our data shows a transcriptional switch during the differentiation of primordia in C. cinerea comprising 11% of the protein-encoding genome being up- or down-regulated. Differential transcription analyses carried out in other basidiomycetous fungi, such as Agrocybe

aegerita, Cordyceps militaris and Ganoderma lucidum, have revealed similar or even more

extensive transcriptional switches [69-71]. Taking the differential expression threshold used

in the present study (log2(fruiting body/vegetative mycelium)≥ 3), 25% and 30% out of

18474 A. aegerita loci are up-regulated in fruiting bodies and vegetative mycelium,

respectively [69]. Similarly, C. militaris up-regulates 40% loci during fruiting body formation

[70], while G. lucidum boosts the transcription of at least 27% measured loci in its sexual

organs [71].

The enrichment among the up-regulated loci in S1P of the functional annotation tag "N-

methyltransferase" (histone-lysine N-methyltransferase, 7 loci in total) suggests that

epigenetic regulation of gene expression or chromosome dynamics may play an important

role during sexual development in C. cinerea. In Saccharomyces cerevisiae, histone H3

lysine 4 trimethylation marks the sites where double-strand breaks, the first events during

Chapter 2 Developmental transcriptomics of C. cinerea

interhomolog recombination, occur [72]. Up-regulation of histone-lysine N-

methyltransferases in S1P is in accordance with karyogamy and early meiotic phases taking place at stage 1 primordia during fruiting body formation [6].

Several previously characterized nematotoxic and insecticidal proteins [18, 20] were found to

be up-regulated in S1P. Among these proteins, lectins appear to be most abundant. The

fruiting body-specific ricin B-like lectins CCL1 and CCL2 were previously shown to be toxic

to C. elegans due to the binding to α1,3-fucosylated N-glycan cores in the intestine of L1

worms [59]. In accordance to a previous study [73], we found that the tetrameric galectin

CGL1 (CC1G_05003) was highly induced in C. cinerea fruiting bodies. Similar to CCL1 and

CCL2, CGL1 and its isogalectin CGL2 (CC1G_05005) showed toxicity against C. elegans

which was dependent on binding to a Galβ1,4Fucα1,6-epitope on N-glycan cores present in

the worm intestine [60]. Protease inhibition is another defense strategy against predation in

C. cinerea fruiting bodies. We found the locus encoding the serine protease inhibitor

Cospin1 (CC1G_09480) and its isoprotein Cospin2 (CC1G_09479) among the top 50 most

highly transcribed and differentially expressed loci in S1P. Cospin1 was previously shown to

be toxic against Drosophila melanogaster larvae indicative of its role in fruiting body defense

against arthropod predation [20]. In this study, we demonstrate that CC1G_11805 is toxic for

C. elegans and A. aegypti larvae, analogous to the toxicity of aerolysin/ETX pore-forming

domain-containing Bacillus sphaericus protein towards larvae of the mosquito Culex

quinquefasciatus [74], suggesting that pore formation is another defense strategy of C.

cinerea primordia against predators. The high absolute transcription of these defense loci in

S1P can be explained by the extensive resource allocation to these organs and the

significance of these organs for reproduction of the fungus. The expression of putative

secreted and antibacterial proteins in C. cinerea vegetative mycelium, on the other hand,

probably reflects the confrontation of this tissue with bacterial competitors although the

antibacterial activity of these proteins has still to be demonstrated. In summary, the

differential expression of defense proteins in the different tissues of C. cinerea is an

Chapter 2 Developmental transcriptomics of C. cinerea adaptation of the fungus to the different environmental challenges with which these tissues are confronted.

Surprisingly, with the exception of cfs1, none of the genes previously shown to play a role in

C. cinerea fruiting body formation were differentially expressed in stage 1 primordia.

Mutations in rmt1, ubc2 and snf5 [14, 15, 17] block development before initials are formed

(48 h before S1P develop). In contrast, dst1, dst2, eln3 and exp1 are involved in processes taking place in stage 2 primordia (24 h after S1P), immature fruiting bodies (48-72 h after

S1P) and decaying fruiting bodies (72 h after S1P) [6, 11, 12, 16]. Thus, a possible explanation of our results is that these genes might be regulated before or after the formation of S1P takes place. Intriguingly, the mutation of cfs1 blocks development right at the transition between initials and S1P. This observed up-regulation of cfs1 in S1P supports a function for this cyclopropane fatty acid synthase in the development of S1P from initials.

The Velvet protein regulon, including genes nsdD, rosA, veA and stuA among others, coordinates sexual development and secondary metabolism in filamentous ascomycetes

[54, 65]. Velvet domains structurally resemble the RHD-like fold present in the transcription factor NF-κβ that plays a central role in animal immunity, suggesting a common evolutionary origin for these two protein families [75]. Overexpression of veA and nsdD in A. nidulans induces the formation of nursing Hülle cells surrounding the cleistothecia [65]. Homologous genes in C. cinerea, CC1G_12219 and CC1G_06391, showed high expression in S1P suggesting a function of these loci in the gene circuitry involved in fruiting body development in this fungus. On the other hand, rosA, an A. nidulans transcription factor inhibiting sexual development in low-carbon culture, is expressed in A. nidulans asexual hyphae where it represses the transcription of sexual development regulators such as nsdD, veA and stuA

[76]. Similarly, rosA-homologous C. cinerea gene CC1G_07059, is expressed in vegetative mycelium where it might be inhibiting sexual development. Taken together, these results

Chapter 2 Developmental transcriptomics of C. cinerea

suggest a conserved role of the Velvet protein regulon during sexual development in the

ascomycete A. nidulans and the basidiomycete C. cinerea.

Genes encoding Velvet domain-containing proteins were also found to be up-regulated in fruiting bodies of S. commune and L. bicolor [51, 52]. However, (with the exception of velB

and kapA) genes encoding Velvet-associated proteins such as RosA/NosA, StuA, NsdD,

PpoA, LaeA, FphA or CryA [54], do not show inter-species conservation of differential

expression. Similar lack of transcriptional conservation between the three analyzed

basidiomycete species was observed for orthologs to transcription factors shown to be

involved in S. commune sexual development [67, 68]. Taken together, this evidence

suggests that basidiomycetes show species-specific divergence of transcriptional regulation

in orthologous genes similar to plants [77].

A broader comparison of the transcriptomes of the three analyzed basidiomycete species,

comprising all the genes differentially expressed in C. cinerea during sexual development,

revealed the presence of a conserved gene regulation circuitry among basidiomycetes

during fruiting body formation. As previously observed in fruiting bodies of ascomycetes [78],

our orthology analysis showed that a large fraction of genes differentially expressed in S1P

corresponds to loci which are not present in the basidiomycetes L. bicolor and S. commune.

These results are in agreement with previous observations in plants and animals that genes

associated with sexual reproduction rapidly evolve [79, 80]. The existence of clusters

comprising conserved up- or down-regulated loci with little inter-species expression

variability, suggests that regulation of these loci evolved under stabilizing selection [81].

Sequence and expression conservation might imply an essential role of these genes in

fruiting body development and sexual reproduction in basidiomycetes. Interestingly, L.

bicolor and C. cinerea orthologs to transcription factors described previously as important for

sexual development in S. commune, such as Bri1, Hom1, Gat1, Fst3, C2h2, Fst4 or Hom2

[67, 68], do not show conserved transcriptional conservation in basidiomycetes, suggesting

Chapter 2 Developmental transcriptomics of C. cinerea

that novel regulatory pathways related to sexual development did evolve once speciation

occurred. As a proof of concept, Traeger and collaborators found that the fruiting body

specific transcription factor pro44 (orthologous to the velvet regulon-associated transcription factor nsdD) in the ascomycetes Sordaria macrospora and Pyronema confluens is a core

regulator of perithecia maturation. S. macrospora deficient in pro44 was shown to be sterile

and unable to produce mature perithecia [78]. In addition to a Velvet domain-containing

protein, the expression of the RNA binding protein mei2 (CC1G_01879), a master meiosis

regulator in yeast and plants [82, 83] protecting meiosis-specific transcripts from degradation by the DSR-Mmi system, was increased in S1P or YFB of C. cinerea, S. commune and L.

bicolor. Similarly, the transcriptional regulation of spo11, encoding a protein inducing meiotic

recombination in S. cerevisiae and C. cinerea [84], was also conserved. This induction of

meiosis regulators in multiple species reflects the role of mushrooms in the production and

dispersal of basidiospores.

With regard to the comparative transcriptome analysis of the three basidiomycete species, it

should be noted that in case of S. commune and L. bicolor, only data of monokaryotic

vegetative mycelia was available [51, 52], whereas in case of C. cinerea, the transcriptome

of an isogenic homodikaryotic mycelium was determined. Thus, the degree of conserved

transcriptional regulation during sexual development between C. cinerea, L. bicolor and S.

commune is potentially larger than observed.

Lastly, the proteome of an organism provides a more direct image of its phenotype than the

transcriptome [85]. Detection of proteins using label-free shotgun mass spectrometry fails to

detect low abundance proteins in complex total extracts and allows only semi-quantitative

estimation of relative protein amounts. These properties are in contrast to the superior

standardization and sensitivity achieved by state of the art nucleotide sequencing

technologies. Nevertheless, LC-MS spectra showed the presence of peptides derived from

the nematotoxic lectin CCL1 (CC1G_11778) and the nematotoxic/insecticidal aerolysin/ETX

Chapter 2 Developmental transcriptomics of C. cinerea

pore-forming domain-containing protein CC1G_11805 in the PBS-soluble protein extract of

C. cinerea stage 1 primordia, indicating that these cytoplasmic toxins are indeed expressed at protein level in these organs.

The recent development of gene targeting tools in C. cinerea [13, 86], will help to test whether some of the genes identified as differentially regulated during sexual development in

multiple basidiomycete species, play a role in this process.

Conclusions

In this work, we show that sexual reproduction in C. cinerea A43mutB43mut involves the

differential transcription of at least 11% of its protein-coding genome. Differentially

transcribed genes include several genes coding for defense proteins that protect fruiting

structures and vegetative mycelia from predators and competitors, respectively. Moreover,

our data infers a role of the Velvet protein family during fruiting body formation in

basidiomycetes and thus, a conserved role of this protein family during sexual development

in dikaryotic filamentous fungi. Finally, the result of the comparative transcriptome analysis

of C. cinerea, S. commune and L. bicolor suggest that a conserved set of orthologous genes regulates sexual development in the phylum Basidiomycota. Additional experiments addressing the function of the differentially expressed gene products are required to confirm these hypotheses.

Availability of supporting data

The data sets supporting the results of this article are available in the ArrayExpress repository under the accession number E-MTAB-1968 (http://www.ebi.ac.uk/arrayexpress/).

Chapter 2 Developmental transcriptomics of C. cinerea

Authors' contributions

DFP prepared samples for RNA-seq and LC-MS, performed the differential expression and

comparative transcriptome analyses and wrote the manuscript. CL carried out the LC-MS

analysis of the PBS-soluble proteins from S1P and VM. NSJVDV cloned CC1G_11805,

recombinantly expressed it in E. coli, and performed the toxicity assays on C. elegans and A.

aegypti. MA and MK were involved in the experiment design and conception as well as in the

data analysis and critical revision of the manuscript. All authors read and approved the final

version of the manuscript.

Acknowledgments

We thank W. Rudin and P. Müller (Swiss Tropical and Public Health Institute, Basel,

Switzerland) and M. O. Hengartner (Institute of Molecular Life Sciences, University of Zürich,

Switzerland) for supplying A. aegypti eggs and C. elegans worms, respectively. We are grateful to scientific staff of the Functional Genomics Center Zurich, in particular to Andrea

Patrignani for the library construction and Michal Okoniewski for his support in the mapping

and counting of the sequenced reads. This project was supported by the Swiss National

Science Foundation Grant 31003A_130671 (to MK) and ETH Zürich Grant ETH-34 11-2 (to

MK).

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Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Chapter 3

Innate immunity in fungi: Transcriptional regulation and target

specificity in the defense response of the model mushroom

Coprinopsis cinerea.

David Fernando Plaza1, Stefanie Sofia Schmieder1, Anna Lipzen2, Erika Lindquist2, Markus

Aebi1, Markus Künzler1.

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

2 Genomic Technologies, Joint Genome Institute, Walnut Creek, California, United States of

America

Contributions:

Sample preparation

RNA extraction and quality control

Differential expression analysis

qRT-PCR validation of gene expression

Antisense transcription analysis

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Innate immunity in fungi: Transcriptional regulation and target

specificity in the defense response of the model mushroom

Coprinopsis cinerea

David Fernando Plaza1, Stefanie Sofia Schmieder1, Anna Lipzen2, Erika Lindquist2,

Markus Aebi1, Markus Künzler1.

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

2 Genomic Technologies, Joint Genome Institute, Walnut Creek, California, United States of

America

Abstract

Coprinopsis cinerea inhabits an environment rich in microbes where it interacts with different antagonists. To study the transcriptional response of this fungus to the presence of fungivorous nematodes, bacterial competitors, mechanical hyphal damage or bacterivorous nematodes, the poly(A)-positive transcriptome of C. cinerea challenged with these stimuli was sequenced. In the presence of the fungivorous nematode A. avenae, C. cinerea was found to specifically induce the expression of previously characterized nematotoxic lectins.

On the other hand, interaction with Gram-positive and Gram-negative bacteria induced the expression of loci encoding different putative lysozymes. These results demonstrate that C. cinerea has the ability to recognize specific predators or nutrient competitors and to mount an appropriate defense response.

Keywords: Coprinopsis cinerea, innate immunity, nematode predation, bacterial competition, nematotoxic lectin, putative lysozyme, defense response specificity, antisense transcription,

RNA-seq.

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Introduction

Interactions between organisms can be either beneficial or detrimental for the partners involved. In order to avoid the detrimental effects caused by antagonists, in particular, multicellular organisms have evolved sophisticated defense mechanisms, comprising strategies to distinguish between self and non-self [1, 2], as well as the production of defense molecules, such as proteins [3-5], RNAs [6] and secondary metabolites [7, 8]. It is thought that such systems originally evolved to prevent the fusion of somatic conspecifics that were genetically different [9, 10]. Cytoplasmic and transmembrane pattern recognition receptors

(PRRs) specifically recognizing conserved microbe- (MAMPs) or damage- (DAMPs) associated molecular patterns have been described and characterized in cnidarians [11], annelids [12], mollusks [13], arthropods [14] and chordates [15]. Plants also recognize

MAMPs and DAMPs using PRRs and share effector mechanisms with animals, including the production of reactive oxygen [16-18] and nitrogen [19, 20] species, as well as the expression of toxic proteins [21], antimicrobial peptides [22, 23] and secondary metabolites

[24]. Conserved signaling pathways have been described in these organisms, usually involving mitogen-activated protein kinases [20, 25], suggesting that the innate immune system is an ancient and widespread trait that appeared very early in evolution. In this regard, fungi are expected to also contain such a system but to date the available information about this system is scarce.

One of the main aspects of immunity is the ability of an organism to distinguish between self and non-self. Fungi are known to recognize non-compatible cells of their own kind as part of their mating system [26, 27] or by a mechanism referred to as vegetative heterokaryon incompatibility (HI) [26, 28]. Latter mechanism, known from the filamentous ascomycetes

Podospora anserina and Neurospora crassa, involves the recognition of non-compatible hyphae via cytoplasmic proteins resembling PRRs and containing HET and STAND domains which leads to an extensive transcriptional response, including the induction of genes encoding for toxins [26, 28]. Little is known about the recognition of other organisms including

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation competitors, pathogens, predators and parasites by fungi and fungal responses to such encounters. In Aspergillus, it was shown that the master regulator of secondary metabolite production LaeA is responsible of its resistance to feeding by larvae of the fruit fly Drosophila melanogaster [29]. In agreement with these results, it was recently shown that challenge of vegetative mycelium with fungivorous collembola induced the formation of fruiting bodies and the synthesis of toxic secondary metabolites, suggesting that Aspergillus is able to recognize its predator and mount an appropriate response [29, 30]. In the same regard, it was shown that A. fumigatus responds to the presence of competing actinomycetous bacteria by producing antibacterial polyketides [31]. It was demonstrated that the response of the fungus depends on direct physical contact between the bacterial and fungal filaments and that the induction of the biosynthetic gene clusters followed the acetylation of histones mediated by

Saga/Ada [32].

We have recently shown that the coprophilous model mushroom Coprinopsis cinerea transcribes a broad array of genes encoding defense proteins in the vegetative mycelium and fruiting bodies that target bacterial competitors and animal predators challenging the respective tissues of this fungus (Plaza et al., in press). In addition, we have demonstrated in previous work that two nematotoxic defense proteins from Coprinopsis, CGL1 and CGL2, were induced in vegetative mycelium challenged with the predatory nematode Aphelenchus avenae [3]; however, the specificity and broadness of this response remained unclear.

In order to resolve these issues, we assessed the genome-wide transcriptional response of the vegetative mycelium of C. cinerea against nematode predation and bacterial co-culture.

The results of this study show that loci encoding nematotoxic and potentially bactericidal proteins are specifically induced in response to nematode predation and bacterial co- cultivation, respectively. In addition, anti-sense transcription in the form of poly(A)-positive transcripts was shown to significantly affect the expression of selected loci in a stimulus- specific manner.

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Methods

Bacteria and nematode cultivation

Bacillus subtilis 168 and Escherichia coli Nissle 1917 were plated from frozen stocks on YMG plates (0.4% yeast extract, 1% malt extract, 50 mM glucose and 1.5% agar) and grown for 18 h at 37°C. Single colonies were inoculated in 2.5 ml YMG (0.4% yeast extract, 1% malt extract, 50 mM glucose) broth and grown to stationary phase at 37°C under constant shaking. Bacteria were then washed twice in sterile-filtered PBS (135 mM NaCl, 2.5 mM KCl,

10 mM Na2HPO4 and 17.5 mM KH2PO4; pH 7.4) and their OD600 was adjusted with PBS to

0.0005 before use. The fungivorous nematodes Aphelenchus avenae (a gift from Richard

Sikora at the University of Bonn) and Bursaphelenchus willibaldi (kindly provided by Dr. Ute

Schönfeldt, Zossen, Germany) were replicated at 20°C on Agaricus bisporus (Hawlik Euro-

Pilzbrut GmbH) or Botrytis cinerea grown on PDA plates (Difco), respectively, and harvested by the Baermann funnel method [33]. Bacterivorous Caenorhabditis elegans N2 was grown on NGM plates (51 mM NaCl, 2.5 g/L bacteriological peptone, 13 mM cholesterol and 1.7% agar) seeded with E. coli OP50 and harvested with PBS. After harvest, all nematodes were transferred to agar plates supplemented with 200 µg/mL G418, 50 µg/mL Nystatin and 100

µg/mL Ampicillin, and incubated for 48 h to eliminate all the residual bacteria or fungi. Before use, nematodes were suspended and washed twice in sterile-filtered PBS and adjusted to a density of 2500 worms/mL

C. cinerea cultivation and challenge

C. cinerea monokaryotic Okayama7 (O7) or dikaryotic A43mutB43mut (AB) mycelia were grown on 30 mL YMG agar plates covered with sterile cellophane discs at 37°C in the dark for 96 h or 72 h, respectively. One of the following organisms was applied in 200 µL sterile- filtered PBS on the mycelium and incubated for 72 h at 24°C in biological triplicates: (I)

Approx. 500 mixed-stage A. avenae; (II) Approx. 500 mixed-stage B. willibaldi (O7 only); (III)

Approx. 500 mixed-stage C. elegans (O7 only); (IV) OD600: 0.0005 B. subtilis 168 (O7 only) or

(V) OD600: 0.0005 E. coli Nissle 1917 (O7 only). To mimic the tissue damage of the hyphae

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation by the fungivorous nematodes, three cellophane-covered YMG plates on which C. cinerea

O7 had grown for 96 h were cut with a sterile scalpel every 24 h during 72 h. As a negative control, 200 µl sterile-filtered PBS was added to three cellophane-covered YMG plates on which C. cinerea O7 or AB had grown for 96 h or 72 h, respectively, in the dark at 37°C.

Mycelia were harvested and flash frozen using liquid nitrogen in individual tubes and stored at -80°C.

Total RNA extraction

Mycelia were lyophilized in a VirTis Freezemobile for 18 h. 20 mg/replicate/treatment lyophilized tissue was lysed in three FastPrep FP120 homogenization steps of 45 s at 4.5,

5.5 and 6.5 m/s in the presence of 250 mg 0.5 mm glass beads, cooling the samples for 5 min on ice between the steps. RNA was extracted using 1 mL Qiazol (Qiagen) and 0.2 mL chloroform (ReagentPlus, Sigma-Aldrich). The mixture was centrifuged at 12000 xg for 15 min at 4°C and RNA from the upper aqueous phase was washed in-column using the

RNeasy Lipid Tissue Mini Kit (Qiagen) and eluted in 60 µL RNase-free water. Concentration and quality of the purified RNA were determined with a Qubit (1.0) fluorometer (Life

Technologies) and a Bioanalyzer 2100 (Agilent), respectively. Samples with a 260/280 nm ratio of 1.8–2.1 and a 28S/18S ratio of 1.5–2 were used for library construction.

Illumina HiSeq 2000 library construction

TruSeq Stranded mRNA Sample Prep Kit (Illumina) was used to construct expression libraries (three biological replicates per treatment). Briefly, total RNA (1 μg) from each biological replicate was poly(A)-enriched and then reverse-transcribed into double-stranded cDNA in the presence of Actinomycin during first-strand synthesis. Double-stranded cDNA was fragmented, end-repaired and adenylated before being ligated to TruSeq adapters and selectively enriched by PCR. Concentration and quality of the enriched libraries were assessed using Qubit (1.0) fluorometer and LabChip GX (PerkinElmer), showing an average fragment size of 260 bp. Libraries were normalized to 10 nM with 10 mM Tris-HCl pH 8.5

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation supplemented with 0.1% Tween 20. In addition, TruSeq PE Cluster Kit v3-cBot-HS (Illumina) was used to generate clusters from 10 pM pooled normalized libraries. Paired-end sequencing was performed on an Illumina HiSeq 2000 at 2 X 101 bp or single end 100 bp using the TruSeq SBS Kit v3-HS (Illumina).

qRT-PCR validation

RNA-seq results were validated by qRT-PCR. Single-stranded cDNA from three (A. avenae challenge or non-challenge control) or two (scalpel-damaged mycelia) biological replicates was synthesized using Transcriptor Universal cDNA Master (Roche) from 2 µg total RNA. 20

µL qRT-PCR reactions were mixed in three technical replicates per primer set and sample, containing 900 nM forward and reverse primers designed to span exon-exon junctions (Table

1), 10 µL 2X FastStart Universal SYBR Green Master (Rox, Roche) and 1 ng/µL cDNA template. qRT-PCR was performed in a Rotor-Gene 3000 (Corbett Life Science) with the following thermal profile: a hold step at 95°C for 15 min followed by 40 cycles of 95°C for 15 s, 62°C for 30 s and 72°C for 30 s. In order to control the specificity of amplification, the reaction was concluded with a melting curve analysis ramping from 55°C to 99°C in steps of

1°C every 5 s. PCR efficiencies and cycle thresholds were obtained using LinRegPCR 12 and differential expression ratios were calculated by the CT difference formula [34]. Tubulin beta chain (CC1G_04743) was used as a house keeping normalizer. In addition, water or 1 ng/µL RNA were included as negative control reactions. To further validate the significance of the RNA-seq-derived differential expression analysis, the constitutive expression of an array of housekeeping loci commonly used in qRT-PCR normalization [35-41] was verified in the sequencing datasets after library size normalization.

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Table 1. Primers used in qRT-PCR validation of gene induction in response to A. avenae predation.

Locus name Primer name Sequence 5'→3' Tm°C CC1G_04743 TubM1Fw GTCATGTCCGGTATCACCAC 62 TubM1Rv GGGAAAGGAACCATGTGGA 61 CC1G_02104 Cc2104FwqRTPCR TGGCAATCCATGACCAGCAGCAATG 70 Cc2104RvqRTPCR AAAGGCTCGCTCGGGCACGAAAC 72 CC1G_02355 Cc2355FwqRTPCR TTTCTGCCCAGCTTGACTAC 62 Cc2355RvqRTPCR CGTGCTTCGAGATCAAATTCGTCG 66 CC1G_05003 CGL1-rtPCR-fwd CCAAGAGGATTAAGGAGAACG 59 CGL1-rtPCR-rev CTAGCAAGCATCCTAAGCG 60 CC1G_05005 CGL2_2FwqRTPCR ACAATGCGGAGAACTCTTTGT 62 CGL2RvqRTPCR CCAGCGAGAATCCTAAGCA 61 CC1G_06684 Cc6684FwqRTPCR TCTCTGGGTTGGCCAGATCATC 65 Cc6684RvqRTPCR TTGGGTGCAGAGAACCAATC 62 CC1G_06698 Cc6698FwqRTPCR CTGAAGTGTACTTCAACTGGCTCG 65 Cc6698RvqRTPCR TTATGCGACCTGTCGTTCGAC 64 CC1G_08593 Cc8593FwqRTPCR TATGACGACCTCGACAGCCGTG 68 Cc8593RvqRTPCR AAACGCTTAGCGCCTCC 62 CC1G_09966 Cc9966FwqRTPCR TGCGACATTGGGTGTTCAG 62 Cc9966RvqRTPCR GGTCCACAGAGAGGATATTGC 61 CC1G_10077 qPCR_77_f GGTAGTAGTCGCCTGAATCG 61 qPCR_77_r CTCCGGTGCAGAGGAATAC 61 CC1G_10726 Cc10726FwqRTPCR TCTTGCGGGTCCTATCGTG 63 Cc10726RvqRTPCR TTAAACAGGAGGGTGCGCG 64 CC1G_11847 Cc11847FwqRTPCR TCTGCATCACCACTTCTGAC 61 Cc11847RvqRTPCR AGGGCAGAGGCCAGTCAAG 66 CC1G_12246 Cc12246FwqRTPCR AACAGCGTCCTCGCGTCACTCAAC 71 Cc12246RvqRTPCR GTCATAGGACTTGCTCATCTGG 62 CC1G_12718 CC12718FwqRTPCR ACAGGTCCCCTATCTCGATGAC 64 CC12718RvqRTPCR TGGTGAGTCCGTAGAATGGCTTG 66 Tm°C: Melting temperature

Bioinformatic analysis

Fastqc files were used to filter and trim the reads to be mapped. Strand-specific reads were mapped to the third annotation (September 2010) of the C. cinerea Okayama7 #130 genome

(C. cinerea Sequencing Project, Broad Institute of Harvard and MIT

(http://www.broadinstitute.org/)) using TopHat. Illumina mapped reads were counted using

Cufflinks and all the sequences were deposited in the ArrayExpress database

(http://www.ebi.ac.uk/arrayexpress/). To determine the percentage of loci showing baseline expression, 5 reads/locus were taken as a minimal threshold. Reads per million mapped reads (RPM) were calculated for every locus in order to scale-normalize all the samples according to library size [42]. Fold change per locus and a Student's t-test comparing the different challenge and control libraries were computed using normalized sense-RPMs.

Volcano plots comparing every treatment with the negative control were constructed. A

Student's t-test p-value ≤ 0.05 (-log10 (p-value) > 1.3) and fold change ≥ 4 were the criteria established to classify a locus as significantly induced in a treatment. A Venn diagram was

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation constructed to identify commonly regulated loci among selected treatments using VENNY

[43]. In addition, a centroid-linkage hierarchical clustering analysis of the computed fold changes was calculated with Cluster 3.0 and heat maps were generated and visualized using

TreeView 3 [44] in order to identify gene clusters including loci specifically induced by each treatment . Functional annotation based on sequence similarity for loci found to be significantly induced was deduced by PSI-BLAST. SignalP 4.1 [45] and TMHMM v. 2.0 [46] were used to predict the presence of secretion signal and transmembrane helices in differentially expressed loci.

Analysis of antisense transcription

Strand specific poly(A)-positive transcriptional data was obtained for every locus in every sample. The fraction of protein-coding transcription in a locus X was calculated by dividing the number of sense reads in locus X by the total number of reads mapping to the same locus (sense reads from locus X / (sense reads from locus X + antisense reads from locus

X)). Genome-wide protein coding transcription profiles were constructed for every sample by ordering all the loci in the genome in descending order according to their fraction of protein coding transcription (sense reads/total reads). The area below the curves was calculated using the trapezoidal rule and the difference in this area for every treatment in comparison to the non-challenged control was statistically tested using a Student's t-test. In addition, a gene-by-gene comparison between the treatments and the negative control was performed.

Genes showing t-test p-values ≤ 0.05 were considered to have a significantl y different fraction of protein-coding transcription compared to the negative control.

Results

General transcriptome features of C. cinerea vegetative mycelium

Illumina RNA-seq stranded libraries were constructed and sequenced in biological triplicates for C. cinerea mycelia exposed to fungivorous nematodes, bacterial co-culture, mechanical damage or the presence of a non-fungivorous (bacterivorous) nematode. Sequencing of 83

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation these libraries showed that 82-86% of the open reading frames (ORF) in the genome of C. cinerea were significantly transcribed (more than 5 reads mapping to the ORF). A total 1.2 billion sense and antisense reads (100 bases each) were mapped to the 37.5 Mb genome of

C. cinerea Okayama-7, accounting for a sequencing output of nearly 120 billion bases (3200 times the genome size) (Table 2).

Table 2. General features of C. cinerea mycelia exposed to different biotic and abiotic stress conditions.

Mapped reads (sense + antisense) Treatment Replicate 1* Replicate 2* Replicate 3* Total Mean % of loci transcribed O7 + PBS 3.31E+07 4.14E+07 4.54E+07 1.20E+08 82.3 AB + PBS 6.20E+07 6.14E+07 5.87E+07 1.82E+08 86.2 O7 + A. avenae 2.51E+07 4.90E+07 3.87E+07 1.13E+08 82.1 AB + A. avenae 6.65E+07 5.92E+07 5.34E+07 1.79E+08 86.2 O7 + B. willibaldi 2.85E+07 5.23E+07 3.53E+07 1.16E+08 82.1 O7 + C. elegans 3.14E+07 5.28E+07 4.31E+07 1.27E+08 82.4 O7 + mechanical damage 2.40E+07 5.14E+07 4.08E+07 1.16E+08 82.6 O7 + E. coli 5.53E+07 4.77E+07 5.09E+07 1.54E+08 82.5 O7 + B. subtilis 4.86E+07 3.78E+07 3.63E+07 1.23E+08 83.3 Total No. mapped reads 1.23E+09 Approximate read length 100 b Approxinate total output 123 Gb * Biological replicates

A. avenae predation specifically induces the expression of loci encoding nematotoxic proteins in C. cinerea.

To study the inducible innate defense response of C. cinerea against nematode predation, C. cinerea strains O7 and AB were co-cultivated with the fungivorous nematodes A. avenae and

B. willibaldi for 72 hours, and their poly(A)-positive transcriptomes were analyzed by RNA- seq. Genes encoding the nematotoxic lectins CGL1 (CC1G_05003), CGL2 (CC1G_05005) and CC1G_10077 were shown to be specifically induced by A. avenae predation (Table 3,

Figure 2A and 2C). Other nematotoxic lectins, such as CCL1 (CC1G_11778) and CCL2

(CC1G_11781), were induced to a lower extent by A. avenae and B. willibaldi, respectively; nevertheless, this induction was not statistically significant (p-values of 0.1 and 0.2, respectively). On the contrary, the bacterivorous nematode C. elegans did not induce the expression of these loci suggesting that the mere presence of live nematodes on the C. cinerea mycelium is not sufficient to trigger the transcription of loci encoding nematotoxic

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation lectins (Figure 2B and 2C, Table 3). In general, defense loci were not induced by B. willibaldi indicating that C. cinerea is capable of distinguishing between different fungivorous nematodes, subsequently triggering a species-specific defense response (Figure 2B).

Fungivorous nematodes feed on the cytoplasmic content in hyphae by piercing a hole in their cell wall and membrane using a specialized feeding apparatus (stomatostylet) which resembles the needle of a syringe [47]. As an attempt to mimic the cell damage in hyphae upon nematode predation, C. cinerea vegetative mycelium was repeatedly injured with a sterile scalpel blade. Expression of nematotoxic lectins was not induced by this treatment

(Figure 2C), demonstrating that cell-wall damage and cytoplasmic leakage are not sufficient to trigger the nematotoxic response observed in C. cinerea exposed to A. avenae predation.

Loci encoding NACHT, HET or HeLo domain-containing proteins which have been shown to be induced as part of the transcriptional program associated to HI in P. anserina [26], were not found to be induced in C. cinerea under any of the treatments applied on the mycelia. To validate these RNA-seq data, the expression of a selection of genes showing statistically significant induction upon A. avenae challenge was confirmed by qRT-PCR (Figure 1). A closer look at the expression of reported housekeeping loci [35-41] in the RNA-seq datasets revealed that none of these genes was up- or down-regulated by any of the treatments applied on Coprinopsis (Table 4).

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Figure 1. qRT-PCR validation reflects the transcriptional behavior observed for several genes by RNA-seq upon A. avenae challenge. The average log2 (fold change) of three

(non-challenged and Aa-challenged) or two (mechanical damage (MD)) biological replicates

(3 PCR reactions/replicate) that were simultaneously sequenced are shown. RNA-seq is displayed as the average of three biological replicates normalized based on the RNA-seq expression of Tubulin B chain. The gene encoding the Glutathione S-transferase

CC1G_12778 was included as a control for constitutive expression. Bars represent the standard deviation.

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Table 3. C. cinerea Okayama7 loci significantly up-regulated in response to every treatment.

Treatment/KEGG ID PSI-Blast Aa/NC Bw/NC Ce/NC MD/NC Ec/NC Bs/NC SignalP TMHMM A. avenae (Aa) CC1G_01501 fruit-body specific protein a 4.1 - - - - - Y 0 CC1G_02104 peroxidase 5.4 - - - - - Y 0 CC1G_02355 hypothetical protein 4.4 - - - - - Y 0 CC1G_02622 LmrCD-specific DARPin 5.2 - - - - - N 0 CC1G_03076 lysozyme 5.2 - - - - - Y 0 CC1G_04169 WSC domain-containing protein 5.2 - - - - - Y 0 CC1G_05003 CGL1 Coprinopsis 5.0 - - - - - N 0 CC1G_05005 CGL2 Coprinopsis 4.1 - - - - - N 0 CC1G_05809 glycosyl hydrolase family 18 5.8 - - - - - Y 0 CC1G_06698 RTA1-like protein 20.4 - - - - - N 7 CC1G_08057 rhsE 4.4 - - - - - N 0 CC1G_08593 hypothetical protein 6.2 - - - - - Y 1 CC1G_09966 nmrA-family protein 4.9 - - - - - N 0 CC1G_10077 Ricin B lectin 4.9 - - - - - N 0 CC1G_10726 hypothetical protein 28.4 - - - - - Y 0 CC1G_11792 hypothetical protein 5.0 - - - - - N 1 CC1G_11847 lysozyme 9.4 - - - - - Y 0 CC1G_12246 ubiquinol cytochrome-c reductase complex 6.8 - - - - - N 7 CC1G_14365 hypothetical protein 6.1 - - - - - N 0 CC1G_14558 hypothetical protein 5.6 - - - - - N 0 B. williballdii (Bw) CC1G_02830 hypothetical protein - 5.9 - - - - N 0 CC1G_03073 hypothetical protein - 7.4 - - - - N 0 CC1G_03493 CsbD domain-containing protein - 5.1 - - - - N 0 CC1G_04080 hypothetical protein - 7.5 - - - - N 0 CC1G_04378 bacteriophage tail tape meausure protein - 4.5 - - - - N 3 CC1G_07359 UDP-N-acetylmuramoylalanine--D-glutamate ligase - 6.6 - - - - N 0 CC1G_07531 hypothetical protein - 4.0 - - - - N 0 CC1G_07533 conidiation-specific protein 6 - 5.2 - - - - N 0 CC1G_07696 HHE domain-containing protein - 4.5 - - - - N 0 CC1G_08790 Ricin B lectin - 5.3 - - - - Y 0 CC1G_10162 ThiJ/PfpI family protein - 4.4 - - - - N 0 CC1G_11449 hypothetical protein - 4.6 - - - - N 0 CC1G_11558 3-oxo-5-alpha-steroid 4-dehydrogenase - 5.3 - - - - N 4 CC1G_14442 hypothetical protein - 4.5 - - - - N 0 CC1G_15585 glutathione-dependent formaldehyde-activating protein - 4.7 - - - - N 0 CC1G_00715 hypothetical protein - 6.4 - - - - N 0 CC1G_01479 hypothetical protein - 5.8 - - - - Y 0 CC1G_02301 hypothetical protein - 6.1 - - - - N 1 CC1G_04294 Sgt1 - 4.9 - - - - N 0 CC1G_04463 ATP-dependent DNA helicase - 4.9 - - - - N 0 CC1G_04669 N-acetyl-gamma-glutamyl-phosphate reductase - 4.6 - - - - N 0 CC1G_08820 putative dead deah box helicase protein - 4.0 - - - - N 0 CC1G_11651 hypothetical protein - 7.8 - - - - N 0 Mechanical damage (MD) CC1G_02583 f5/8 type C domain protein - - - 6.0 - - N 1 CC1G_03098 P-loop containing nucleoside triphosphate hydrolase protein - - - 4.8 - - N 0 CC1G_03514 hypothetical protein - - - 4.3 - - N 0 CC1G_09359 hypothetical protein - - - 4.9 - - N 0 CC1G_10157 hypothetical protein - - - 5.0 - - N 0 CC1G_11387 cyclopropane-fatty-acyl-phospholipid synthase - - - 4.5 - - N 0 CC1G_11620 nipped-B-like protein - - - 4.6 - - N 0 CC1G_12367 hypothetical protein - - - 5.7 - - N 7 CC1G_15356 hypothetical protein - - - 5.9 - - N 0 E. coli (Ec) CC1G_00122 cytochrome P450 - - - - 4.1 - N 0 CC1G_01525 TAL1 - - - - 4.5 - N 0 CC1G_02052 YCII-related domain protein - - - - 5.6 - N 0 CC1G_02062 N-alpha-acetyltransferase 60-like - - - - 9.4 - N 0 CC1G_02166 hypothetical - - - - 9.3 - N 1 CC1G_02345 malate synthase - - - - 5.7 - N 0 CC1G_02382 lipolytic enzyme - - - - 5.4 - Y 0 CC1G_02862 snoal-like polyketide cyclase family protein - - - - 6.5 - Y 0 CC1G_02908 alcohol dehydrogenase - - - - 4.8 - N 0 CC1G_03442 endonuclease/exonuclease/phosphatase - - - - 5.1 - Y 0 CC1G_03541 hypothetical protein - - - - 16.7 - Y 0 CC1G_04927 hypothetical protein - - - - 5.9 - N 3 CC1G_05515 D-amino-acid oxidase - - - - 5.5 - Y 0 CC1G_05607 alginate regulatory protein AlgP - - - - 5.1 - Y 0 CC1G_05864 lolT-1 Coprinopsis 0.0, PLP-dependent transferase - - - - 7.1 - N 0 CC1G_05914 ammonium transporter - - - - 32.0 - N 11 CC1G_06488 urea transporter - - - - 5.5 - N 15 CC1G_06620 isocitrate lyase - - - - 4.5 - N 0 CC1G_06972 ndb1 (nad(p)h dehydrogenase b1) - - - - 4.1 - N 0 CC1G_07061 hypothetical protein - - - - 4.8 - N 0

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

CC1G_07630 phosphatidylinositol 4-kinase - - - - 4.1 - N 3 CC1G_07735 succinate:fumarate antiporter - - - - 5.0 - N 0 CC1G_08094 high affinity methionine permease - - - - 6.0 - N 12 CC1G_08394 hypothetical protein - - - - 8.5 - N 0 CC1G_08758 hypothetical protein - - - - 4.3 - Y 0 CC1G_08822 WSC domain-containing protein - - - - 4.1 - Y 0 CC1G_08888 acyl-CoA carboxylate CoA-transferase - - - - 9.1 - N 0 CC1G_11310 priA protein - - - - 5.6 - Y 1 CC1G_11374 PLAC8-domain-containing protein - - - - 4.3 - N 0 CC1G_11786 hypothetical protein - - - - 6.1 - N 4 CC1G_12758 acyl-CoA N-acyltransferase - - - - 20.3 - N 0 CC1G_12964 citrate synthase - - - - 4.1 - N 0 CC1G_12996 Ricin B lectin - - - - 4.3 - Y 0 CC1G_13124 ammonium transporter - - - - 7.9 - N 9 CC1G_13213 glycerophosphoryl diester phosphodiesterase - - - - 12.2 - Y 0 CC1G_13246 hypothetical protein - - - - 6.2 - N 5 CC1G_13826 hypothetical protein - - - - 7.4 - N 4 CC1G_14125 mitochondrial carrier - - - - 5.1 - N 3 CC1G_14829 Glutathione Transferase Gte1 - - - - 8.4 - N 0 CC1G_15187 hypothetical protein - - - - 4.0 - N 0 CC1G_15681 hypothetical protein - - - - 8.3 - N 0 CC1G_15703 cytochrome p450 - - - - 4.7 - N 0 CC1G_08300 hydrophobin-251 - - - - 10.5 - N 1 B. subtilis (Bs) CC1G_03042 lysozyme - - - - - 5.6 Y 0 CC1G_05798 fruit-body specific gene A - - - - - 4.2 Y 0 CC1G_09605 hypothetical protein - - - - - 8.9 N 0 CC1G_10004 hypothetical protein - - - - - 5.1 Y 0 CC1G_08311 monocarboxylate permease - - - - - 4.7 N 11 Ec+Bs CC1G_00718 hypothetical protein - - - - 6.6 6.4 Y 1 CC1G_05600 hypothetical protein - - - - 10.9 11.7 Y 0 CC1G_08056 rhsE - - - - 20.6 15.9 Y 0 CC1G_08433 hypothetical protein - - - - 42.8 11.1 N 0 CC1G_09365 triacylglycerol lipase - - - - 6.8 11.5 Y 0 CC1G_14477 lysozyme - - - - 6.9 6.2 N 0 Aa+Ec CC1G_02441 hypothetical protein 6.6 - - - 5.2 - Y 0 CC1G_04734 Med17 domain-containing protein 4.5 - - - 5.7 - Y 0 CC1G_07582 hypothetical protein 8.9 - - - 5.1 - Y 0 CC1G_09529 hypothetical protein 5.0 - - - 5.7 - Y 0 CC1G_10384 O-methylsterigmatocystin oxidoreductase 4.6 - - - 6.7 - N 0 CC1G_06684 LysM domain-containing protein 4.2 - - - 15.4 - Y 0 C. elegans (Ce)+Ec CC1G_02581 hypothetical protein - - 8.0 - 31.1 - Y 2 Bw+Ec CC1G_03339 beta-Ig-H3/Fasciclin - 9.6 - - 4.8 - Y 0 Bw+Ce CC1G_04017 hypothetical protein - 4.9 5.8 - - - N 0 Aa+Bs CC1G_05472 rhsE 9.0 - - - - 6.7 Y 0 CC1G_03047 lysozyme 7.0 - - - - 4.8 N 0 Ce+Bs CC1G_08818 hypothetical protein - - 17.7 - - 38.1 Y 0 CC1G_13803 hypothetical protein - - 7.2 - - 12.7 N 0 Aa+Ce+Ec CC1G_13818 MFS general substrate transporter 9.1 - 5.4 - 16.1 - Y 9 MD+Bs CC1G_15139 metalloprotease - - - 4.0 13.4 Y 0 Ce+Ec+Bs CC1G_01042 PAP2 superfamily protein - - 21.7 - 42.9 35.0 Y 0 CC1G_05219 KapM protein - - 7.5 - 14.5 9.3 Y 0 As thresholds of significant differential expression, fold (treatment/negative control)≥ 4 and student's t-test p-value ≤ 0.05 (from three biological replicates per treatment) were used

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Table 4. Housekeeping genes do not show up- or down-regulation in C. cinerea mycelia subjected to different culture conditions

log2(fold) Locus Functional annotation ABAa/NC O7Aa/NC O7Bw/NC O7Ce/NC O7MD/NC O7Ec/NC O7Bs/NC Reference CC1G_00758 Beta tubulin -0.07 -0.28 0.32 0.15 0.03 0.13 0.34 Wan H 2011 CC1G_00876 Tubulin beta chain 0.11 -0.51 0.48 0.08 -0.07 -0.08 0.04 Wan H 2011 CC1G_03523 Histone H2B -0.05 -0.25 0.04 0.17 0.65 0.57 -0.03 Ferreira 2012 CC1G_03676 Histone H2B 0.02 -0.26 -0.02 0.19 0.32 0.58 -0.08 Ferreira 2012 CC1G_03927 Actin 0.35 -0.48 -0.44 -0.07 -0.04 0.02 0.07 Huggett 2005 CC1G_04355 Actin 0.01 -0.28 0.18 0.11 0.00 0.00 0.16 Huggett 2005 CC1G_04743 GADPH 0.06 -0.32 -0.58 0.00 0.03 -0.03 -0.06 Huggett 2005 CC1G_06184 GADPH 0.11 -0.60 -0.76 0.03 -0.04 -0.02 -0.12 Huggett 2005 CC1G_07639 Ubiquitin -0.08 -0.21 -0.02 0.20 0.49 0.11 -0.23 Silveira 2009 CC1G_08232 Ubiquitin 0.27 -0.40 -0.42 -0.08 -0.13 -0.02 -0.16 Silveira 2009 CC1G_09116 Ubiquitin C 0.52 -0.51 -0.25 -0.21 -0.28 0.00 -0.24 Silveira 2009 CC1G_09117 Cyclophilin -0.38 0.49 1.12 0.07 1.13 1.52 1.37 Langnaese 2008 CC1G_09572 Ribosomal protein S27a 0.19 -0.66 -0.20 -0.11 -0.31 0.38 0.07 de Oliveira 2011 CC1G_11833 Ribosomal protein L19 0.06 -0.20 0.06 0.10 0.18 -0.17 -0.41 de Oliveira 2011 CC1G_13048 Ribosomal protein L11 -0.27 -0.48 -0.71 0.25 0.25 1.26 0.09 de Oliveira 2011 CC1G_13649 Ribosomal protein L32 -0.01 -0.26 0.36 0.12 0.05 -0.05 0.12 de Oliveira 2011 CC1G_15352 Hsp90 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Aursnes 2011

C. cinerea O7 induces the expression of lysozymes in the presence of bacteria and nematodes.

To evaluate the transcriptional response of C. cinerea vegetative mycelium to the presence of Gram-positive and Gram-negative bacteria, hyphae were co-cultivated in triplicates with B. subtilis 168 and E. coli Nissle 1917, respectively. In the presence of B. subtilis, 28 loci were found to be differentially expressed with 18 and 10 loci being up- or down-regulated, respectively (Figure 2D, and Tables 3 and 5). In comparison, 81 loci were differentially expressed in C. cinerea in the presence of E. coli, representing 60 and 21 significantly induced or repressed loci, respectively (Figure 2D, and Tables 3 and 5). A comparison of the different sets of genes induced by each treatment revealed that CC1G_14477, a locus encoding a predicted lysozyme, was commonly induced by B. subtilis and E. coli, indicating that C. cinerea can sense the presence of conserved molecular patterns in Gram-positive and Gram-negative bacteria inducing the expression of putative antibacterial genes (Table

3). In addition the expression of the lysozyme-encoding loci CC1G_03042 and CC1G_03047 was significantly increased by the presence of B. subtilis (the latter being also induced by A. avenae) in the C. cinerea culture. Intriguingly, hierarchical clustering also shows that

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

CC1G_03076 is induced by A. avenae, E. coli, B. subtilis, and scalpel damage; suggesting that up-regulation of this lysozyme might be part of a general stress response in C. cinerea.

Figure 2. A. avenae predation and bacterial co-cultivation induce the expression of nematotoxic lectins and putative bactericidal proteins in C. cinerea vegetative mycelium. (A) Volcano plots showing a genome-wide differential expression analysis of C. cinerea O7 co-cultivated in three biological replicates with A. avenae (O7Aa), B. willibaldi

(O7Bw), C. elegans N2 (O7Ce), E. coli Nissle 1917 (O7Ec) or B. subtilis 168 (O7Bs). To partially mimic the mechanical damage inflicted in hyphae by fungivore feeding, three biological replicates were also cut with a scalpel (O7MD) and their transcriptomes sequenced. Genes showing log2 (treatment/NC) higher than 2 or less than -2, and -log10

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

(Student's t-test-derived p-values calculated from three biological replicates) > 1.3 were considered to be significantly up-regulated (red boxes) or down-regulated (blue boxes). (B)

Venn's diagram computed for genes significantly up-regulated by fungivore feeding (O7Aa and O7Bw), the presence of a bacterivorous nematode (O7Ce) or scalpel-inflicted hyphal damage. C. cinerea O7 triggers a nematode-specific defense response that is not mimicked by the damage with scalpel. (C) Centroid linkage hierarchical clustering analysis calculated on the mean fold changes for every treatment compared to the negative control shows the presence of two loci clusters specifically induced by C. cinerea O7 and AB in response to A. avenae predation (O7Aa and ABAa). The three induced clusters include loci encoding nematotoxic lectins (marked in red). (D) Clusters specifically induced by co-culture with bacteria. E. coli and B. subtilis trigger the expression of the lysozyme CC1G_03076 in C. cinerea O7. A statistical testing of the gene expression for the biological replicates sequenced shows that lysozyme CC1G_03047 is significantly induced by the presence of A. avenae or B. subtilis (Table 3). Pearson correlation coefficients (PC) are shown for some gene clusters.

Potential PRRs in C. cinerea are down-regulated in the presence of A. avenae or E. coli.

Down-regulation of loci caused by the set of treatments applied on C. cinerea was also analyzed. Most of the genes modulated by the different culture conditions (26 out of 48) were found to encode hypothetical proteins with no predicted functional annotation. Interestingly, potential PRRs containing LRR (CC1G_08259) or WD40 (CC1G_06907) domains were also suppressed by the presence of the fungivorous nematode A. avenae or the Gram-negative bacterium E. coli Nissle 1917, suggesting that these two organisms might modulate the host response in Coprinopsis by decreasing its microbe-recognition capabilities (Table 5).

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Table 5. C. cinerea Okayama7 loci significantly down-regulated in response to every treatment.

Tretament/KEGG ID PSI-Blast NC/Aa NC/Bw NC/Ce NC/MD NC/Ec NC/Bw SignalP TMHMM A. avenae (Aa) CC1G_09526 endoglucanase-4 4.1 - - - - - Y 0 CC1G_09644 alpha-galactosidase 5.8 - - - - - N 0 CC1G_13605 hypothetical protein 20.5 - - - - - N 0 CC1G_13985 hypothetical protein 6.7 - - - - - N 0 CC1G_14464 reverse transcriptase/ribonuclease H 6.2 - - - - - N 0 B. willibaldi (Bw) CC1G_03086 hypothetical protein - 7.9 - - - - Y 0 CC1G_06494 C2H2 finger domain-containing protein - 6.1 - - - - N 0 CC1G_09242 SNF2 superfamily protein - 28.2 - - - - N 0 CC1G_10994 hypothetical protein - 4.4 - - - - N 2 CC1G_11779 hypothetical protein - 9.0 - - - - N 0 CC1G_14460 hypothetical protein - 4.6 - - - - N 0 CC1G_14846 CNH-domain-containing protein - 4.8 - - - - N 0 CC1G_15718 hypothetical protein - 13.4 - - - - N 0 CC1G_15756 WSC domain-containing protein - 10.2 - - - - Y 0 C. elegans (Ce) CC1G_02106 hypothetical protein - - 6.7 - - - Y 0 CC1G_09458 extracellular tungstate binding - - 4.1 - - - N 0 CC1G_13558 hypothetical protein - - 5.1 - - - Y 0 CC1G_15088 hypothetical protein - - 5.4 - - - N 0 Mechanical damage (MD) CC1G_08269 hypothetical protein - - - 4.6 - - Y 0 CC1G_08983 hypothetical protein - - - 6.7 - - N 7 E. coli (Ec) CC1G_01035 hypothetical protein - - - - 8.8 - N 0 CC1G_01577 glycosyl hydrolase family 62 protein - - - - 6.7 - Y 0 CC1G_01879 RNA recognition motif 2 partial - - - - 4.1 - N 0 CC1G_02999 hypothetical protein - - - - 4.2 - N 0 CC1G_06017 tyrosinase central domain-containing protein - - - - 5.3 - Y 0 CC1G_08259 Leucine Rich Repeat domain protein - - - - 16.5 - N 0 CC1G_10148 hypothetical protein - - - - 4.1 - N 0 CC1G_10470 tyrosinase central domain-containing protein - - - - 8.0 - Y 0 CC1G_10494 hypothetical protein - - - - 4.7 - N 5 CC1G_11188 ycaC protein - - - - 8.1 - N 2 CC1G_11580 hypothetical protein - - - - 4.5 - N 0 CC1G_12408 hypothetical protein - - - - 4.2 - N 0 CC1G_12509 galactose mutarotase-like protein - - - - 4.4 - N 0 CC1G_14013 cytochrome P450 - - - - 4.5 - N 1 CC1G_14014 O-methylsterigmatocystin oxidoreductase - - - - 4.6 - N 0 CC1G_14164 alphaN-acetylglucosamine transferase - - - - 4.8 - N 1 B. subtilis (Bs) CC1G_01487 hypothetical protein - - - - - 6.9 N 0 CC1G_04915 hypothetical protein - - - - - 9.0 N 4 CC1G_05337 hypothetical protein - - - - - 6.4 N 7 CC1G_05968 hypothetical protein - - - - - 5.6 N 1 CC1G_08393 hypothetical protein - - - - - 6.2 Y 1 CC1G_14873 hypothetical protein - - - - - 4.4 N 0 Ec+Bs CC1G_03158 hypothetical protein - - - - 5.9 7.4 Y 3 CC1G_09799 other/AgaK1 protein kinase - - - - 5.9 7.1 N 0 CC1G_10006 tyrosinase - - - - 4.3 4.1 N 0 Aa+Ec CC1G_06907 WD40 domain-containing protein 5.8 - - - 6.8 - N 1 CC1G_15644 hypothetical protein 10.2 - - - 13.6 - Y 0 Aa+Bs CC1G_15616 extracellular tungstate binding 4.9 - - - - 6.2 N 0 As thresholds of significant differential expression, fold (negative control/treatment) ≥ 4 and student's t-test p-value ≤ 0.05 (from three biological replicates per treatment) were used.

Treatment-specific regulation of antisense transcription in C. cinerea

The construction of stranded poly(A)-positive cDNA libraries allowed us to study the effect of different interaction setups on the production of locus-specific anti-sense transcripts. The fraction of sense reads over the total sum of reads mapping to every annotated locus in the 92

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

C. cinerea O7 genome was calculated. A genome-wide profile for antisense transcription was constructed for every sample and biological replicate, and the amount of antisense transcription per sample was indirectly measured by the trapezoidal rule as the area under the curve in the profile. The three resultant areas for every treatment were compared to the areas calculated in the negative control to determine whether the interaction with nematodes or bacteria, as well as the scalpel-caused damage in the hyphae, could trigger a detectable increase or decrease in the total amount of antisense transcripts produced. A minor area increase reflecting a general decrease in the proportion of antisense transcription relative to the total amount of transcription was observed when B. willibaldi or B. subtilis were co- cultivated with C. cinerea; however the difference was not statistically significant (Figure 3A).

On the contrary, individual gene comparisons between the different treatments and the negative control revealed that the ratio of protein-coding RNA relative to the total amount of

RNA mapping to some loci was significantly different (Figure 3B). For some loci, a general correspondence between the extent of their protein-coding relative to total transcription, and their differential expression was observed. Genes showing significant increases or decreases in the protein-coding relative to total transcription ratio as measured in the A. avenae- challenged mycelium and the non-challenge control were also up- or down-regulated upon A. avenae predation, respectively (Figure 3C).

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Figure 3. Stress and microbial interaction significantly affect the amount of antisense transcription for some loci in C. cinerea Okayama-7. (A) The fraction of protein coding transcription (sense reads) per locus was calculated for every sample sequenced. Loci were ordered in descending order according to the amount of protein-coding relative to the total amount of transcription (increasing fraction of antisense transcription) and the area under the curves was measured. The area under the curve of every treatment was compared to the non-challenged control using a Student's T-test. Genome-wide, none of the treatments induced a statistically significant change in the amount of poly(A)-positive antisense transcription observed. (B) A gene-by-gene comparison between the different treatments and 94

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation the non-challenged control shows that every treatment significantly (p-value < 0.05) affects the amount of protein-coding relative to the total transcription for some loci. (C) C. cinerea O7 loci showing a significant change in the fraction of protein-coding transcription after being challenged with the fungivorous nematode A. avenae. Orange and blue bars show the fraction of sense relative to total transcription in non-challenged and A. avenae-challenged C. cinerea O7, respectively. Grey bars represent the transcription log2 (fold) between A. avenae-challenged and non-challenged C. cinerea O7. Dashed lines mark the differential expression thresholds set for this study (log2 (fold) > 2 and log2 (fold) < -2). Stars represent the p-values calculated for the amount of antisense transcription in O7Aa and O7NC *:

0.01

(though not significant) induction of the gene and vice versa.

Discussion

Similar to plants [48], fungi are non-motile organism lacking an adaptive immune system and specialized immune cells. Nevertheless, plants and fungi have evolved inducible innate immune systems which allow them to repel predators [3, 29, 49]. Whereas the plant innate immune system is rather well characterized, the fungal one is largely unknown.

Previous reports suggested that both filamentous asco- and basidiomycetes are able to induce specific responses to biotic stresses [3, 29, 32]; nevertheless, the extent of this response on genome-wide level was unknown. Our results show that C. cinerea has the ability to discriminate between different stimuli and respond to these stimuli by the induction of specific gene sets. We show that C. cinerea induces the production of cytoplasmic toxic lectins or secreted lysozymes when challenged with the fungivorous nematode A. avenae or bacteria, respectively, demonstrating that this basidiomycete not only distinguishes between different non-self stimuli but also responds by inducing appropriate defense proteins.

Evidence suggests that B. willibaldi is able to feed on the ascomycete B. cinerea but not on

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

Coprinopsis (Stefanie Schmieder, personal communication), indicating that the induction of a specific defense response against fungivorous nematodes other than A. avenae should not be discarded.

Similar changes in gene expression have been observed during HI reactions between ascomycetes like Podospora and Neurospora [26, 28]. These reactions were shown to be triggered by ligand binding to cytoplasmic WD40 domain-containing proteins and are proposed to have originally evolved as a recognition mechanism to detect pathogens [50].

Even though genes encoding HET and STAND domain-containing proteins were not found to be differentially expressed after any of the treatments applied, C. cinerea induced the expression of carbohydrate binding proteins as a defense response to nematodes as it was shown for P. anserina in the course of HI, indicating that similar regulatory pathways could be involved in HI and defense.

Cytoplasmic lectins CGL1, CGL2 and CC1G_10077 [Stefanie Schmieder, personal communication] were specifically induced by nematode predation in C. cinerea. These lectins have been shown to be toxic to the bacterivorous nematode C. elegans when administered recombinantly in IPTG-induced E. coli BL21 [51]. In addition, the expression of CGL1 and

CGL2 was previously established to be increased when C. cinerea was challenged for 72 h with A. avenae [3]. The use of lectins as defense effectors of the innate immune system is widely spread among distantly related taxa. In the mammalian intestine, blood group antigen- binding galectins, such as Gal-4 and Gal-8, kill blood group antigen B-decorated bacteria such as E. coli O86 [52]. On the other hand, the human lectin Gal-3 induces cell death in the pathogen Candida albicans mediated by binding to β-1,2-linked oligomannans on the outer most layer of the fungal cell wall [53]. In plant immunity, lectins related to GNA [54], Hevein

[55], Nictaba [56] or Ricin [57], as well as legume lectins [58] and Jacalins [59], have been shown to be insecticidal and induced upon insect feeding [21, 60]. Accordingly, fungal lectins have also been described to play a role in the defense of sexual organs, mainly based on

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation their toxicity towards and on their specificities for glycans of insects, nematodes and amoeba

[3, 51, 61, 62].

The capacity of lectins to recognize non-self glycans [3] makes them promising PRR candidates too. For example, dectins are receptors expressed on the surface of antigen- presenting cells sensing mannosylated epitopes from fungi. Dectins 2 and 3 form heterodimers recognizing surface alpha-mannans on C. albicans and inducing activation of

NF-κB [63]. Dectins belong to the family of C-type lectin-like receptors containing cytoplasmic immunoreceptor tyrosine-based activation motifs (ITAM). Similarly, galectins have also been described to function as PRRs binding non-self glycans and inducing immunity [64]. As mentioned above, cytoplasmic fungal lectins are specific to non-fungal glycans and represent an interesting group of PRRs that would engage in recognition of non-self-glycan epitopes and, at the same time, act as an effector.

Intriguingly, genes encoding RTA1 and the hypothetical protein CC1G_10726 were the most highly induced by A. avenae predation. RTA1 is a membrane transporter found to confer resistance to the inhibitor of yeast growth 7-aminocholesterol [65, 66]; however, there are no reports on the synthesis of aminocholesterol derivatives in nematodes that would allow us to draw a connection between predation and the induction of RTA1 in Coprinopsis.

Several lysozymes were found to be induced upon interaction of C. cinerea with fungivorous nematodes and bacteria. Interestingly, the lysozyme CC1G_03042 was specifically induced by the Gram-positive bacterium B. subtilis 168. On the contrary, the lysozyme CC1G_03076 was un-specifically induced by most of the treatments (though only significantly in mycelia challenged with A. avenae), suggesting that this gene is part of a general stress response program in C. cinerea. Intriguingly, the lysozyme CC1G_14477 was significantly induced in the presence of B. subtilis and the Gram-negative bacterium E. coli. Lysozymes are enzymes hydrolyzing the β-1,4-glycosidic bond linking monomers of N-acetylmuramic acid and N-

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation acetylglucosamine in the bacterial peptidoglycan [67]. Although the presence of an outer membrane reduces the accessibility of lysozymes to the cell-wall of Gram-negative bacteria, lysozyme induction has been observed in coelomocytes from the earthworm Eisenia Andrei exposed to E. coli [68]. Furthermore, the C-type lysozyme MgCLYZ from the mussel Mytilus galloprovincialis was shown to be have lytic activity against the Gram-negative bacteria

Vibrio anguillarum, Enterobacter cloacae, Pseudomonas putida, Proteus mirabilis and B. aquimaris [69], indicating that some lysozymes are part of the innate defense response against Gram-negative microorganisms. The specificities of fungal lysozymes are largely unknown and a thorough biochemical analyses of these enzymes is needed.

We found extensive antisense transcription in Coprinopsis, with a big proportion of protein- coding loci showing anti-sense reads being mapped to them. Moreover, antisense transcription of some specific loci was significantly influenced by environmental stimuli.

Antisense transcripts are known to co- and post-transcriptionally regulate gene expression in fungi in a number of ways [70]. Firstly, antisense-transcribing RNA polymerase II can interfere with the synthesis of RNA taking place in the sense direction of the same locus by direct polymerase-polymerase collision [71]. Secondly, cis antisense transcription recruits chromatin-remodeling enzymes leading to silencing of the neighboring protein-coding transcription [72]. Thirdly, antisense-mediated intron retention can be caused by antisense transcripts annealing to complementary intron splicing sites, interfering in this way with the splicing machinery [73]. Lastly, duplexes composed by sense and antisense transcripts can be degraded by RNA interference endonucleases in the cytoplasm [74]. Similarly, CAGE analysis of different human cell types revealed the presence of up to 313003 promoters driving the expression of cis antisense transcripts, demonstrating that antisense transcription is widespread in human cells [75]. C. cinerea encodes homologs to the main components of the RNA-mediated silencing machinery, such as Dicer, argonaute and RdRP. Furthermore, gene silencing mediated by the expression of homologous hairpin RNAs has been demonstrated for Coprinopsis [76], showing that posttranscriptional regulation might play a

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation role in the biology of this basidiomycete; nevertheless, additional biological evidence is needed to validate the relevance of antisense transcription in the regulation of gene expression of C. cinerea.

In conclusion, RNA-seq of C. cinerea challenged with fungivorous nematodes and bacteria showed that the induction of genes encoding nematotoxic lectins and lysozymes is part of the transcriptional response of C. cinerea against predators and bacterial competitors. The specificity of this response suggests that Coprinopsis has the ability to discriminate between different kinds of non-self stimuli and to regulate its gene expression accordingly. Finally, widespread antisense transcription affecting different genes in a stimulus-specific manner was observed in C. cinerea hyphae. This study provides transcriptional evidence on the existence of a tightly regulated innate immune system in C. cinerea; supporting previous reports showing the structural and functional characterization of defense effectors in this fungus [3, 51, 61].

Acknowledgments

This project was supported by the Swiss National Science Foundation Grant

31003A_130671. Illumina libraries from C. cinerea Okayama-7 were sequenced as part of the DOE Joint Genome Institute's Community Sequencing Program 'Functional genomics in the model mushroom Coprinopsis cinerea'. The work conducted by the U.S. Department of

Energy Joint Genome Institute is supported by the Office of Science of the U.S. Department of Energy under Contract No. DE-AC02-05CH11231. We are also thankful to the scientific staff at the Functional Genomics Center Zurich, in particular to Catharine Aquino for the library construction and Michal Okoniewski for his support with the mapping and counting of the sequenced reads corresponding to the C. cinerea A43mutB43mut samples.

Chapter 3 Innate immunity in C. cinerea: Transcriptional regulation

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Chapter 4

Cloning, expression and partial functional characterization of

putative defense proteins of Coprinopsis cinerea

David F. Plaza1, Esther Ketelaars1, Tina Dimler1, Markus Künzler1, Markus Aebi1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

Contributions:

Protein cloning in E. coli and P. pastoris

Protein expression and purification

C. elegans toxicity assays

Subtilisin inhibition assay

Lactosyl-sepharose affinity chromatography

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Cloning, expression and partial functional characterization of

putative defense proteins of Coprinopsis cinerea

David F. Plaza1, Esther Ketelaars1, Tina Dimler1, Markus Künzler1, Markus Aebi1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

Abstract

Fungi produce effector molecules, e.g. proteins and secondary metabolites to defend themselves against predators, parasites and competitors. To functionally characterize putative defense proteins from the model mushroom Coprinopsis cinerea, two putative protease inhibitors (CC1G_05299 and CCIA1/CC1G_06934), a predicted pore-forming toxin

(Caerolysin/CC1G_10318) and the Ricin B fold domain-containing protein CC1G_10603, were cloned and recombinantly expressed in the bacterium Escherichia coli or the yeast

Pichia pastoris. The aspartate protease inhibitor CC1G_05299 was expressed in a soluble form in E. coli BL21 but was neither toxic to Caenorhabditis elegans pmk-1 nor to Aedes aegypti larvae. CCIA1, in contrast, was expressed in E. coli BL21 in soluble form and shown to inhibit C. elegans N2 larval development at the L4 stage. Moreover, purified CCIA1 was demonstrated to inhibit the serine protease Subtilisin Carlsberg in a concentration-dependent manner. For CC1G_10318, a wide range of cloning, expression and purification strategies were used in order to obtain soluble Coprinopsis aerolysin (Caerolysin). None led to the production of soluble Caerolysin, though. Purification of His-tagged Caerolysin from E. coli inclusion bodies was attempted and a small amount of soluble protein was recovered after re-folding. This protein lacked hemolytic activity against erythrocytes from goat, sheep, cow, horse or pig at a 10 µg/mL concentration, however. Similarly, the secreted Ricin B fold domain-containing protein CC1G_10603 was found to be insoluble when expressed in P. pastoris and the project was stopped at this stage. In summary, it appears that the selection of suitable expression and purification systems are major challenges for the biochemical

112

Chapter 4 Characterization of putative defense proteins of C. cinerea characterization of pore-forming proteins and secreted Ricin B fold domain-containing proteins from fungi.

Keywords: E. coli, P. pastoris, putative defense protein, protease inhibitor, Ricin B-fold domain, nematotoxicity.

Introduction

Fungi and plants are non-motile organisms that rely on chemical strategies, i.e. the production of secondary metabolites and protein toxins, to defend themselves against antagonists. In previous chapters, we showed that the expression of defense proteins is, on the one hand, tissue-specific: the defense proteins of the respective tissues (vegetative mycelium or fruiting body) match the type of antagonists these tissues are confronted with.

On the other hand, some defense proteins were shown to be induced upon predation by the fungivorous nematode Aphelenchus avenae. Characterized fungal defense proteins against predators comprise biotin binding proteins [1], protease inhibitors [2], lectins [3-7] and pore forming proteins [8-11]. With the exception of proteins binding biotin with high affinity, C. cinerea encodes [12] several proteins falling into these categories in its genome which are potential defense molecules in the warfare against predators.

Some defense proteins act by starving predators or pathogens from crucial nutrients necessary for their development and survival [13, 14]. Evidence has shown that the biotin- binding proteins Tamavidin 1 and 2 from the basidiomycete Pleurotus cornucopiae are highly toxic against Caenorhabditis elegans, Drosophila melanogaster and Acanthamoeba sp.

Exogenous addition of biotin inhibits Tamavidin toxicity, indicating that this class of protein depletes the growing medium from this vital nutrient [1].

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Protease inhibitors (PIs) are peptides or small molecules which can hinder the activity of proteolytic enzymes. According to sequence homology, PIs can be classified in at least 93 families [15, 16]. Serpins, for example, inhibit serine proteases and represent the largest family known of PIs. Structurally, the members of this PI superfamily contain a Serpin scaffold and a reactive center loop that occupies and covalently binds to the protease catalytic site [17]. Sometimes, proteases are regulated by adjacent domains called propeptides which facilitate protease folding and can temporarily block the catalytic site of the enzyme before activation [18, 19]. Inhibitors such as Pleurotus ostreatus proteinase inhibitor A 1 (POIA1) [20] and Bombyx cysteine proteinase inhibitor (BCPI) [21] structurally resemble the propeptides found in serine or cysteine peptidases. On the contrary, β-trefoil domains are largely versatile in their function and can be found not only in inhibitors such as the Kunitz-soybean trypsin inhibitor (STI) [22] and the papaya protease inhibitor (PPI) [23]; but also in lectins such as C. cinerea lectins 1 and 2 and Clitocybe nebularis lectin CNL [4], among others. PIs have been shown to play a role in the defense of plants and fungi against foraging. Cospin1, for example, is a serine PI highly expressed in the fruiting bodies of C. cinerea that is toxic against larvae of the fruit fly Drosophila melanogaster [2]. In the plant

Solanum nigrum, at least four serine protease inhibitors have been shown to deter herbivorous insects by inhibiting specific intestinal proteases; nevertheless, a dual role in defense and development has not been discarded yet [24]. Some PIs are regulated in response to environmental stimuli. For instance, wound-inducible Pin-II protease inhibitors are induced in response to stress in Capsicum annuum [25]. In this study, two protease inhibitors were cloned, expressed and functionally characterized. Firstly, C. cinerea protease inhibitor A 1 (CCIA1) is homologous to POIA1 and is an interesting defense candidate in C. cinerea. Secondly, CC1G_05299 was initially considered to be a Ricin B-fold lectin; however,

Sabotič and collaborators have recently found that this protein is an aspartate PI that inhibits

Pepsin as well as endogenous aspartate peptidases in C. cinerea (personal communication).

In addition, tissue-specific transcriptome analysis in C. cinerea revealed that CC1G_05299 and its paralog CC1G_05298 were highly expressed in vegetative mycelium and stage 1

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Chapter 4 Characterization of putative defense proteins of C. cinerea primordia, respectively (Plaza et al, 2014; in press), making them promising tissue-specific defense protein candidates.

Lectins are proteins that recognize carbohydrates in a non-catalytic and reversible manner, showing high heterogeneity in structure and ligand specificity [26]. Lectin production has been found to be a crucial component of the defense response of fungi [3-7] and plants [27-

30] against predators and foragers. In fungi, lectins of diverse carbohydrate specificities are developmentally regulated and expressed in the fruiting bodies as defense effectors of these organs. Interestingly, many fungal lectins are not secreted and are specific to non-self glycans [3, 26]. Evidence suggests that fungal lectins recognize carbohydrates exposed in the predator's intestinal lumen and cause toxicity by an unknown mechanism [6, 7]. In C. cinerea, at least three nematotoxic lectins (Coprinopsis galectin 1 and 2, and CC1G_10077) were found to be significantly and specifically up-regulated upon predation by the fungivorous nematode A. avenae. CC1G_10077, C. cinerea lectin 1 and 2 (CCL1 and CCL2) are three fungus-derived Ricin B-fold-containing lectins shown to be toxic against predatory nematodes. Among 23 Ricin B-fold domain-containing proteins Coprinopsis encodes in its genome [12], CC1G_10603 is a secreted polypeptide that was selected for cloning, expression and functional characterization due to its sequence homology to nematotoxic lectins such as CCL1 and CCL2.

Aeromonas hydrophila aerolysin [31] and Laetiporus sulphureus lectin (LSL) [8] belong to the superfamily of beta-pore-forming toxins, playing roles in bacterial virulence and fungal defense, respectively. Beta-barrel pore-forming toxins are secreted as monomers and experience dramatic conformational changes to form oligomeric transmembrane channels when in close proximity to their target cell [32]. Some members of this superfamily, such as

LSL, contain a glycan targeting domain contiguous to the toxin domain [8]. C. cinerea encodes at least three cytoplasmic proteins containing Aerolysin-like domains in its genome

(CC1G_10318, CC1G_08369 and CC1G_11805) [12]. CC1G_10318 and CC1G_11805 have

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Chapter 4 Characterization of putative defense proteins of C. cinerea been previously shown to be highly expressed in sexual organs (Plaza et al, 2014; in press), making them attractive defense candidates against fungivorous nematodes. In order to functionally characterize the pore-forming protein CC1G_10318 too, we cloned and recombinantly expressed it in E. coli.

In this chapter, trials to demonstrate the function of some fungal proteins in the defense against predators and parasites are shown. The studied proteins were selected based on their sequence similarity to previously characterized defense proteins (CCIA1, CC1G_10603,

CC1G_05299 and CC1G_10318) as well as on their tissue specific pattern of expression

(CC1G_05299 and CC1G_10318).

Methods

RNA extraction and cDNA synthesis

Total RNA was extracted from 20 mg lyophilized vegetative mycelium or stage 1 primordia from C. cinerea A43mutB43mut. In brief, mycelium or primordia were lysed with 200 mg 0.5 mm glass beads in 2 mL screw cap tubes in three 45 s steps of FastPrep FP120 homogenization at 4.5, 5.5 and 6.5 m/s, cooling the extracts on ice between steps.

Thereafter, 1 mL QIAzol (QIAGEN) was added and vortexed for 2 min to fully disrupt the hyphae. 200 µL chloroform were added and the extracts were vortexed for 15 s and centrifuged at 12000 xg for 15 min at 4°C. The aqueous phases were mixed in a 1:1 ratio with 70% ethanol and the RNA contained in them was washed and eluted in 60 µL water using RNeasy Lipid Tissue Mini kit columns according to the manufacturer's instructions

(QIAGEN). RNA quality was assessed on a Bioanalyzer Nano chip. Transcriptor Universal cDNA Master (Roche) was used to synthesize cDNA from 2 µg total RNA. cDNA was stored at -20°C for later use.

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CC1G_05299 cloning, expression and purification

A pET24-derived plasmid for the expression of N-terminus-His-tagged CC1G_05299 was constructed using NdeI and NotI restriction sites. cDNA derived from C. cinerea mycelium was used as a template to amplify the spliced transcript encoding the protein. Pfu polymerase in combination with the primers CC1G_05299fw8HisNdeI and

CC1G_05299RvNotI were used to amplify a His-tagged version of CC1G_05299 (Table 1).

3' overhanging adenines were added using Taq polymerase and dATP. The PCR product was inserted into pGEM-T-easy vector (Promega) that was then used to transform chemocompetent E. coli DH5α. Plasmid was extracted with NucleoSpin Plasmid

(MACHEREY-NAGEL) from at least 5 clones selected on a LB plate containing 80 µg/mL X-

Gal, 0.5 mM Isopropyl β-D-1-thiogalactopyranoside (IPTG) and 100 µg/mL ampicillin, and sequenced using the vector primers M13-forward and M13-reverse (Table 1). A plasmid displaying a mutation-free insertion was digested with NdeI and NotI in Buffer O (Thermo

Scientific) and ligated into pre-digested pET24 backbone. CC1G_05299-coding insert and pET24-backbone for ligation were purified from gel using NucleoSpin Gel and PCR Clean-up

(MACHEREY-NAGEL). An aliquot of chemocompetent E. coli DH5α was transformed with the pET24-NHCC1G_05299 ligation product and 3 clones were selected on LB plates supplemented with 50 µg/mL kanamycin. Plasmids were extracted from these clones as previously described and sequenced using the vector primers T7Fwd and T7term (Table 1).

Chemocompetent E. coli BL21 aliquots were transformed with vectors showing no mutations and selected on LB-Kan plates.

Table 1. Cloning and sequencing primers.

Primer name Sequence 5' → 3' Tm°C CC1G_05299fw8HisNdeI GGGGCATATGCATCATCATCATCATCATCACCACGAGCCTGGACGATACAG 79 CC1G_05299RvNotI CGCGGCCGCTCAAGCCTCTTCAACAACCC 78 CCIA1FwNde1 GGGGCATATGTCGACCGGAAAGTTCATTGT 72 CCIA1Fw8HisNde1 GGGGCATATGCACCACCACCACCACCACCACCACTCGACCGGAAAGTTCATTGT 84 CCIA1 rev (BamHI) GGGGGGGATCCTCATTGAGTGGTAACAACA 72 Caerolysinfw2NdeI GGGGCATATGGGTGTTGGCCTCCGATG 74 Caerolfw2His8NdeI GGGGCATATGCACCACCACCACCACCACCACCACGGTGTTGGCCTCCGATG 86 CaerolRvstopNotI CGCGGCCGCTCAGCACGCCTTCTTGAGCTTG 80 CaerolRvHis8NotI CGCGGCCGCTCAGTGGTGGTGGTGGTGGTGGTGGTGGCACGCCTTCTTGAGCTTG 88 CaerolRvlec1stopNotI CGCGGCCGCTTAGCGCTCGACGGTGATCTTGTTCC 81 CC1G_10603FwEcoRI GAATTCAAAATGGGGCTCAGCGCTCTCTACC 72 CC_10603RvHisNotI1 GCGGCCGCTTAATGATGATGATGATGATGATGATGCAGGGTCCATACCTG 79 M13 TGTAAAACGACGGCCAG 59 M13r CAGGAAACAGCTATGACC 56 T7Fwd TAATACGACTCACTATAGGG 54 T7term TGCTAGTTATTGCTCAGCGG 59

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Chapter 4 Characterization of putative defense proteins of C. cinerea

His-tagged CC1G_05299 protein expression was induced with 1 mM IPTG in an OD600: 0.5

BL21/pET24- NHCC1G_05299 culture during 18 h at 24°C. Cells were harvested and disrupted using a French press in PBS supplemented with 1 mM phenylmethanesulfonylfluoride (PMSF). Whole cell extract was centrifuged at 26000 xg to remove cell debris and insoluble proteins. The supernatant was loaded on 1 mL pre- equilibrated Ni-NTA agarose during 1 h at 4°C on a rotating wheel, and the flow-through was discarded. The column was washed with 50 mL PBS containing 10 mM imidazole and pure

His-tagged CC1G_05299 was eluted in 10 mL PBS supplemented with 200 mM imidazole.

Imidazole was removed in a PD-10 desalting column (GE Life Sciences) with a final PBS elution step. Finally, His-tagged CC1G_05299 was concentrated in an Amicon Ultra-4

Centrifugal Filter Unit (3000 NMWL, Merck Millipore) and its concentration measured by

BCA.

Lactosyl-sepharose-mediated affinity chromatography of CC1G_05299

150 µL lactosyl-sepharose were transferred to a Mobicol column and equilibrated with 3 mL

PBS (6 steps of 500 µL each followed by centrifugation at 3000 xg for 2 min). 300 µL His- tagged CC1G_05299 (250 µg/mL) were loaded on the lactosyl-sepharose and incubated on a rotating wheel for 1 h. The column was centrifuged at 3000 xg for 2 min and the flow- through was collected. To remove unbound CC1G_05299, the column was washed with 3 mL PBS. Bound protein was eluted with PBS containing 200 mM lactose in 2 steps of 500 µL each, incubating every time for 10 min at 4°C in a rotating wheel. From all the fractions collected (load, flow-through, wash and elution), 25 µL were mixed in a 1:1 proportion with

4X Laemmli buffer and loaded on two 15% SDS-PAGEs. The first gel was Coomassie- stained and the second used for sequential detection with mouse anti-His and HRP-coupled goat anti-mouse IgG.

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Chapter 4 Characterization of putative defense proteins of C. cinerea

C. elegans toxicity assay

Toxicity of CC1G_05299 and CCIA1 recombinantly expressed in E. coli BL21 towards C. elegans was evaluated as previously described [33]. C. elegans PMK1 and C. elegans N2 were grown on NGM plates (50 mM NaCl, 2.5 g/L bacteriological peptone, 13 mM cholesterol and 1.7% agar) pre-seeded with E. coli OP50 at 24°C until the major proportion of worms corresponded to gravid hermaphrodites and most of the bacteria were consumed. Worms were harvested in deionized and distilled water (ddH2O) and incubated in a solution containing 0.5 N NaOH and 1% NaClO for 10 min. The released eggs were washed three times in a 15 mL conical bottom tube with 8 mL ddH2O and transferred to a 1.5% agar plate to hatch overnight. Larvae were suspended in PBS, counted and their concentration adjusted to 1500 larvae/mL. In a 96 well plate, 20 µL larvae suspension were mixed with 180 µL E. coli BL21 (OD600: 2) previously induced for protein expression with 1 mM IPTG during 18 h at

24°C. As a negative control, an E. coli BL21 strain carrying an empty pET24 construct was used. Positive control corresponded to worms fed with Coprinopsis galectin 2 (CGL2)- expressing E. coli BL21 as previously shown [3]. Four replicates per treatment were used and a Dunn's multiple comparison test was applied to assess the statistical significance in the differences between treatments and controls.

Aedes aegypti toxicity assay

Toxicity of His-tagged CC1G_05299 against A. aegypti Rockefeller (kindly provided by W.

Rudin and P. Müller) larvae was evaluated as described by Künzler and collaborators [33].

600-800 eggs were hatched in a petri dish containing 200 mL ddH2O and 30 mg ground fish food for 20 h at 28°C in the dark. Larvae were transferred to fresh 800 mL ddH2O supplemented with 50 mg ground fish food and incubated for 10 h at 28°C in order to obtain synchronized L2 larvae. Ten L2 larvae/replicate (4 replicates/treatment) were starved in 100 mL fresh ddH2O in 100 mL Schott flasks for 6 h at 28°C before adding 1 mL OD600: 20 IPTG- pre-induced E. coli BL21 (empty vector pET24, pET24-CGL2 or pET24-CC1G_05299NHis).

Larvae were incubated for 96 h at 28°C in the dark and the surviving individuals were

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Chapter 4 Characterization of putative defense proteins of C. cinerea counted. Percentages of surviving larvae/treatment were calculated. A Dunn's multiple comparison test between the empty vector pET24 control and the different treatments was run to test the statistical significance of the results observed.

Cloning, expression and purification of CCIA1

CCIA1 was cloned and expressed as previously described for CC1G_05299 with small modifications. To recombinantly express and purify a His-tagged (N-terminus) version of

CCIA1, the primers CCIA1Fw8HisNde1 and CCIA1 rev (BamHI) (Table 1) were used to amplify CC1G_06934 from stage 1 primordia-derived cDNA. Restriction digestions leading to vector construction were carried out with NdeI and BamHI in 2X Tango Buffer (Thermo

Scientific) at 37°C. Clone selection, as well as protein expression and purification were performed as described for CC1G_05299. In addition, a vector encoding a non-tagged version of CCIA1 was constructed in order to test the toxicity of CCIA1 toward C. elegans N2 when expressed in E. coli BL21. Primers CCIA1FwNde1 and CCIA1 rev (BamHI) (Table 1) were used in the construction of the latter vector.

Subtilisin inhibition assay

1 µM Bacillus licheniformis protease (Subtilisin Carlsberg, Sigma-Aldrich) was preincubated in triplicate with different concentrations of CCIA1 (0, 875, 1750, 2500, 5000 and 10000 nM) for 15 min at 22°C in a flat bottom 96 well plate. 100 µM protease substrate Suc-Ala-Ala-Pro-

Phe-pNA (Bachem) in PBS were added and incubated for 15 min at 22°C.

OD405 was quantified as a measure of substrate degradation.

Expression of the His-tagged Ricin B fold domain-containing protein CC1G_10603 in the P. pastoris system

Protein expression and solubility of CC1G_10603 were assessed according to the manufacturer instructions. The fragment encoding CC1G_10603 tagged with a C-terminus

Histidine peptide and bearing the native secretion signal, was amplified from C. cinerea

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Chapter 4 Characterization of putative defense proteins of C. cinerea stage 1 primordia-derived cDNA and inserted in the expression vector pPICZA (Life

Technologies) with EcoRI and NotI. P. pastoris was then transformed by electroporation with this construct. Expression was induced in BMMH (100 mM potassium phosphate, pH 6.0;

1.34% yeast nitrogen base, 4 x 10–5% biotin and 1% methanol) broth for 48 h at 28°C by adding 1% methanol to the P. pastoris pPICZ-CC1G_10603CH stationary phase culture every 24 h. 500 µL culture samples were collected at time points 0 h, 24 h and 48 h, spinning down the cells and storing separately pellets and supernatants at -20 °C. To evaluate protein solubility, pellets were thawed at 30°C and put on ice. Cells were disrupted using 100 µL

Pichia breaking buffer (50 mM sodium phosphate, pH 7.4; 1 mM PMSF, 1 mM EDTA, 5% glycerol and 1X cOmplete Protease Inhibitor Cocktail) in combination with 0.5 mm acid washed glass beads repeating 8 times one step of vortexing for 30 s followed by 30 s of cooling on ice. Extracts were centrifuged at 14000 xg and 4°C for 10 min, and the supernatants corresponding to the soluble protein fractions were collected. The fractions corresponding to the whole protein extract, soluble protein extract and the culture supernatant were mixed with Laemmli buffer and loaded on two 12% SDS-PAGEs. One of the gels was stained with Coomassie while the other was transferred to a nitrocellulose membrane and incubated with mouse anti-His (IgG1, Qiagen) and HRP-coupled goat anti- mouse IgG (LabForce AG) as primary and secondary antibodies for detection, respectively.

Cloning and expression of Caerolysin and Caerolysin putative lectin domain in E. coli.

His-tagged constructs of Caerolysin and the putative N-terminus lectin domain from this protein were cloned as previously described for CC1G_05299 with small modifications.

Primers Caerolfw2His8NdeI and CaerolRvstopNotI (Table 1) were used to amplify and clone a His-tagged (N-terminus) version of Caerolysin from stage 1 primordia-derived cDNA. A His- tagged (C-terminus) version was cloned with the primers Caerolysinfw2NdeI and

CaerolRvHis8NotI (Table S1). Finally, Caerolfw2His8NdeI and CaerolRvlec1stopNotI (Table

1) were used to amplify the putative lectin domain of Caerolysin. Expression was induced with 1 mM IPTG in OD600: 0.5 cultures as for CC1G_05299; however, expression was carried

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Chapter 4 Characterization of putative defense proteins of C. cinerea out at 24°C and 30°C to assess solubility differences. Expression and solubility were evaluated by comparing whole cell extracts and soluble fractions of induced and non-induced cultures with Coomassie staining and western blotting using mouse anti-His.

In order to improve the solubility of Caerolysin, E. coli ArcticExpress (Agilent Technologies) was transformed with pET24-Caerolysin (a construct encoding the N-terminus-His-tagged version of the protein) and clones were selected on LB plates supplemented with 20 μg/mL gentamycin and 50 μg/mL kanamycin. Expression was induced according to the manufacturer instructions. In brief, two clones were grown overnight at 37°C in LB broth containing 20 μg/mL gentamycin and 50 μg/mL kanamycin. 60 μL cell suspension from each clone were inoculated in 3 mL LB broth without selection antibiotics and grown at 30°C for 3 h before being transferred to 10°C for 10 min under constant shaking. 1 mM IPTG (final concentration) was added and the cultures were grown at 10°C with shaking for 24 h.

Samples were taken at every stage and mixed with Laemmli buffer to be later analyzed by western blot with mouse anti-His.

Purification of Caerolysin from E. coli inclusion-bodies

Caerolysin was purified from inclusion bodies as previously reported [34] with small modifications. Protein expression was induced with 1 mM IPTG in OD600: 0.5 E. coli BL21 transformed with pET24-CaerolysinNHis, and incubated for 18 h at 23°C under constant shaking. Cells were harvested, suspended in lysis buffer (100 mM Tris·Cl, pH 7.0, 5 mM

EDTA, 2 mM β-mercaptoethanol, 5 mM benzamidine·HCl and 1 mM PMSF) and disrupted in a French press. Lysed cell suspension was clarified by centrifugation for 1 h at 22000 xg and

4°C. The supernatant was discarded and the pellet (Approx. 2.5 g) was washed in 10 mL wash buffer (100 mM Tris·Cl, pH 7.0, 5 mM EDTA, 2 mM β-mercaptoethanol, 2 M urea and

2% (w/v) Triton X-100) before being spinned-down for 30 min at 22000 xg and 4°C.

Thereafter, supernatant was discarded and the washing step was repeated twice. The pellet was then suspended in wash buffer without urea and Triton X-100 and centrifuged again.

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Supernatant was discarded and the pellet was suspended with constant steering for 18 h at

4°C in 10 mL extraction buffer (50 mM Tris·Cl, pH 7.0, 5 mM EDTA, 8 M guanidine·HCl and

2 mM β-mercaptoethanol). Next, the solution was centrifuged for 1 h at 100000 xg and 4°C.

The supernatant was recovered and 10 mL ice-cold extraction buffer without guanidine·HCl were added. The protein solution was loaded on 1 mL Ni-NTA agarose that had been previously equilibrated with extraction buffer containing 4 M guanidine·HCl (equilibration buffer), and incubated for 1 h at 4°C under gentle mixing. Flow-through was discarded and the column was washed with 50 mL equilibration buffer. Thereafter, Aerolysin was refolded in column with a gradient of decreasing concentrations of guanidine·HCl (3 mL/step; 3.5 M → 3

M → 2.5 M → 2 M → 1.5 M → 1 M) in 50 mM Tris·Cl, pH 7.0 and 2 mM β-mercaptoethanol.

The partially refolded protein was eluted in a 10 mL solution containing 200 mM imidazole,

50 mM Tris·Cl, pH 7.0 and 2 mM β-mercaptoethanol. Solution volume was reduced to 2.5 mL using an Amicon Ultra-15 Centrifugal Filter Unit (10000 NMWL, Merck Millipore) and imidazole was removed in a PD-10 desalting column (GE Life Sciences). In order to remove the remnants of β-mercaptoethanol, Caerolysin was dialyzed in three steps against 400 mL

PBS pH 7.4 in a Slide-A-Lyzer Dialysis Cassette (10 kDa MWCO, Thermo Scientific).

Concentration was measured by BCA. Protein purity and lack of degradation were assessed by western blot using a mouse anti-His antibody and HRP-coupled goat anti-mouse IgG.

Caerolysin hemolysis assay

1.5 mL heparinized blood from goat, sheep, cow, horse and pig was washed three times with

12 mL PBS pH 7.4 and the resulting cell pellet was suspended in 1.5 mL PBS. 50 µL erythrocytes were mixed with either 50 µL 20 µg/mL Caerolysin or PBS (negative control) in

1.5 mL microcentrifuge tubes and incubated for 30 min at 24°C. Tubes were centrifuged at

2400 xg for 10 min and the supernatants (approx. 100 µL) were split in two wells in a 96 well plate and the optical density at 405 nm was measured with an ELISA reader.

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Results

The Ricin B fold aspartate protease inhibitor CC1G_05299 does not bind lactosyl- sepharose

Using recombinant protein produced from a construct provided by us (see below), Sabotic and collaborators have shown that CC1G_05299 inhibits endogenous aspartate proteases in

C. cinerea (personal communication). Furthermore, they have structural evidence showing that CC1G_05299 is highly similar to the lactose-specific Macrolepiota procera ricin B-like lectin (MPL). In order to test for an potential dual function of CC1G_05299 as protease inhibitor and lectin, we assessed the capacity of CC1G_05299 to bind lactose (Galβ1-4Glc).

For this purpose, a His-tagged version of CC1G_05299 was expressed and purified in E. coli

BL21 (Figure 1).

Figure 1. Soluble His-tagged CC1G_05299 can be expressed in E. coli BL21.

CC1G_05299 was amplified from C. cinerea mycelium-derived cDNA using a forward primer encoding an 8xHis-tag. The resulting product was inserted into the expression vector pET24 with the restriction enzymes NdeI and NotI, and replicated in the E. coli strain DH5α. Plasmid was purified, sequenced and transformed in the expression strain E. coli BL21. Expression was induced for 18 h with 1 mM IPTG and the cells were lysed in a French press. Soluble 124

Chapter 4 Characterization of putative defense proteins of C. cinerea proteins were recovered in the supernatant after centrifugation at 26000 xg and His-tagged

CC1G_05299 was bound to a Ni-NTA column followed by elution with 200 mM imidazole.

Protein was concentrated in a 10 kDa centrifugal filter unit and the imidazole was removed by desalting on a PD-10 column. Fractions corresponding to the whole protein extract

(WCE), soluble protein extract loaded on the Ni-NTA column (Input), input flow-through (FT), wash flow-through (Wash) and Elution were collected, mixed with Laemmli buffer and loaded on a 16% SDS-PAGE.

The purified recombinant protein was loaded onto a lactosyl-sepharose column. The column was washed and the bound protein was eluted using 200 mM lactose. All CC1G_05299 protein was found either in the flowthrough or in the wash fraction and no protein was found in the eluate (Figure 2). These results show that CC1G_05299 does not bind lactose.

Figure 2. CC1G_05299 is not a lactose-binding lectin. 75 µg His-tagged CC1G_05299 from a 250 µg/mL solution were loaded on 150 µL lactosyl-sepharose equilibrated with PBS in a Mobicol column, washed and eluted with 200 mM lactose. Input, flow-through, wash and elution fractions were collected, mixed with Laemmli buffer and loaded in two 15% SDS-

PAGEs. One of the gels was Coomassie-stained whereas the other was transferred to a nitrocellulose membrane and stained with mouse anti-His (1:2000 dilution) and HRP-coupled

125

Chapter 4 Characterization of putative defense proteins of C. cinerea goat anti-mouse IgG. Relative concentration of the collected fractions are also shown.

CC1G_05299 was not found to bind lactosyl-sepharose.

CC1G_05299 is not toxic to C. elegans or A. aegypti larvae

Given the fact that protease inhibitors were shown to be toxic against nematodes (e. g. the L. bicolor homolog of Clitocypin towards C. elegans, Aurélie Deveau, personal communication), and insects [2], we assessed the toxicity of CC1G_05299 towards larvae of C. elegans

PMK1 or A. aegypti Rockefeller as previously reported [33]. Insect and nematode larvae were fed with E. coli BL21 expressing CGL2 or His-tagged (N-terminus) CC1G_05299 in four replicates. Animals were independently fed with bacteria bearing an empty pET24 construct and induced overnight with 1 mM IPTG as a negative control of toxicity. No inhibition of development of insect or nematode larvae by CC1G_05299 was observed (Figure 3)

Figure 3. CC1G_05299 is not toxic to C. elegans or A. aegypti larvae. Toxicity of

CC1G_05299-expressing E. coli towards C. elegans PMK1 (A) and A. aegypti (B) larvae was assessed as previously described [33]. E. coli BL21 expressing the toxic lectin CGL2 was used as a positive control, whereas bacteria carrying empty pET24 vector were used as a negative control of toxicity. CC1G_05299 was not toxic to insect or nematode larvae. Dunn's

126

Chapter 4 Characterization of putative defense proteins of C. cinerea multiple comparison test was used to assess the statistical significance of the differences observed between treatments and the negative control. *: 0.01 ≤ p-value ≤ 0.05

CCIA1 is toxic to C. elegans PMK1 and inhibits the B. licheniformis serine protease

Subtilisin

CC1G_06934 (CCIA1), a C. cinerea protein that is highly similar to POIA1, was cloned, recombinantly expressed as His-tagged protein in E. coli BL21 and purified over Ni-NTA

(Figure 4A). To test the potential toxicity of CCIA1 against nematodes, L1 larvae of C. elegans N2 were fed with CCIA1-expressing E. coli. Worms were found to arrest development at the L4 stage (Figure 4B) suggesting that the target of CCIA1 is important during late C. elegans development. Finally, CCIA1 was tested for inhibition of the activity of

B. licheniformis Subtilisin towards the synthetic substrate Suc-Ala-Ala-Pro-Phe-pNA. CC1A1 was found to inhibit the serine protease Subtilisin in a concentration-dependent manner

(Figure 4C), showing maximum inhibition at concentrations above 5 µM (at least 5:1 inhibitor to protease molar ratio).

Figure 4. CCIA1 is a serine protease inhibitor that stops C. elegans development at L4 stage. His-tagged CCIA1 was purified over a Ni-NTA column (A). CCIA1 was expressed in

E. coli BL21 and soluble protein extracts were prepared (Input) and loaded on a Ni-NTA resin. Flow-through (FT) was collected and the column was washed with PBS (Wash). Pure

CCIA1 was eluted with 200 mM imidazole (Elution). (B) C. elegans N2 develops to L4 larvae but not adulthood when fed with E. coli expressing non-tagged CCIA1. Synchronized L1 larvae were mixed with overnight-IPTG-pre-induced OD600: 2 E. coli BL21 expressing CCIA1

127

Chapter 4 Characterization of putative defense proteins of C. cinerea in four replicates in a 96 well plate. The proportion of worms reaching each larval stage was calculated for every well after 48. The mean proportion of four replicates is shown. Bacteria expressing the nematotoxic lectin CGL2 were fed to C. elegans as a positive control of toxicity. The negative control corresponded to worms fed with E. coli BL21 transformed with an empty pET24 vector (EV). Bars represent the standard deviation calculated for four replicates. Dunn's multiple comparison test was used to assess the statistical significance of the differences observed between treatments and the negative control. *: 0.01≤ p -value ≤

0.05. (C) CCIA1 is a Serine protease inhibitor. 1 µM serine protease from Bacillus licheniformis (Subtilisin Carlsberg) was pre-incubated in triplicate with different concentrations of His-tagged CCIA1 before mixing with the protease substrate Suc-Ala-Ala-

Pro-Phe-pNA in a flat bottom 96 well plate. OD405 was quantified as a measure of substrate degradation. A concentration higher than 5 µM of CCIA1 was necessary to fully inhibit

Subtilisin (corresponding to at least a 5:1 inhibitor to protease molar ratio).

CC1G_10603 is a secreted Ricin B fold domain-containing protein that is insoluble in

P. pastoris.

The secreted Ricin B fold domain-containing protein CC1G_10603 of C. cinerea was cloned into the Pichia expression vector pPICZA in a construct carrying the native secretion signal and a C-terminus His-tag. Protein expression was induced with 1% methanol for 48 h adding new methanol every 24 h and collecting culture samples before methanol addition. Even though CC1G_10603 expression was detected by western blot, no soluble protein was observed in the soluble fraction from the whole cell extracts nor in the culture supernatant

(Figure 5).

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Chapter 4 Characterization of putative defense proteins of C. cinerea

Figure 5. The Ricin B fold domain-containing protein CC1G_10603 is insoluble when recombinantly expressed in P. pastoris. CC1G_10603 was amplified by PCR from C. cinerea stage 1 primordia-derived cDNA and introduced in the Pichia expression vector pPICZA. P. pastoris was transformed by electroporation and clones selected on PDA plates supplemented with Zeocin. Expression was induced on stationary phase liquid cultures by adding 1% methanol (final concentration) every 48 h (two methanol pulses in total, one every

24 h) and culture samples were collected right before every methanol pulse and at the end of the cultivation. Samples were separated into supernatant (SUPER) and cells. Whole cell extracts (WCE) were prepared by mechanical disruption using glass beads and the soluble protein fraction (SF) corresponded to the supernatant from the high speed centrifugation of the WCE. The three fractions from every time point were mixed with Laemmli buffer and loaded on two 12% SDS-PAGEs. The first gel was Coomassie-stained (A) whereas the second was transferred to a nitrocellulose membrane and sequentially detected with mouse anti-His IgG and HRP-coupled goat anti-mouse IgG (B). Coomassie staining shows the induction of the alcohol oxidase AOX1 (approx. 75 kDa) after 24 h of culture in the presence of 1% methanol; nevertheless, insoluble expression of His-tagged CC1G_10603 could be observed by western blot only.

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Chapter 4 Characterization of putative defense proteins of C. cinerea

A wide range of cloning and expression strategies are ineffective to produce soluble

Caerolysin.

CC1G_10318 (Caerolysin) is a C. cinerea protein shown to be highly expressed in stage 1 primordia and displaying sequence similarity to Aeromonas hydrophila aerolysin and

Laetiporus sulphureus lectin (LSL). Different His-tagged constructs corresponding to the full protein or its N-terminus lectin domain were produced in the expression vector pET24 and expressed in E. coli BL21; however, none of them was suitable for soluble expression

(Figures 6 and 7). In addition, a full version of His-tagged Caerolysin was expressed in E. coli ArcticExpress in which it turned to be insoluble too (Figure 8).

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Chapter 4 Characterization of putative defense proteins of C. cinerea

Figure 6. His-tagged CC1G_10318 constructs are insoluble in E. coli BL21 when expressed at different temperatures. Expression of Caerolysin CC1G_10318 tagged with an N- (A) or a C- (B) terminal His8-tag was induced with 1 mM IPTG in E. coli BL21 at 24°C or 30°C. Whole cell extracts (WCE) and soluble fractions (SF) were prepared for induced and un-induced bacteria and loaded on 12% SDS-PAGEs that were transferred to nitrocellulose

membranes and developed with mouse anti-his and HRP-coupled goat anti-mouse IgG.

Figure 7. A His-tagged version of the N-terminus lectin domain from Caerolysin

CC1G_10318 is insoluble in E. coli BL21. A His-tagged clone of the N-terminus region from CC1G_10318 potentially encoding a lectin domain was cloned and expressed in E. coli

BL21 at 24°C and 30°C. Two 14% SDS-PAGEs were loaded with whole cell extracts (WC) and soluble fractions from induced and non-induced bacteria. The first gel was stained with

Coomassie dye (A) while the second was transferred to a nitrocellulose membrane and analyzed by western blot (B). The N-terminus domain of the Caerolysin CC1G_10318 is insoluble when expressed in E. coli BL21.

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Chapter 4 Characterization of putative defense proteins of C. cinerea

Figure 8. CC1G_10318 carrying an N-terminus His-tag is insoluble in E. coli

ArcticExpress. In order to improve the solubility of CC1G_10318, a His-tagged construct of the protein (N-terminus) was expressed in E. coli ArcticExpress (DE3) expressing the cold- adapted chaperonins Cpn10 and Cpn60. Two independent clones selected on a gentamycin/kanamycin LB plate were induced with 0.5 mM IPTG at 10°C for 24 h. The protein was shown to be insoluble in E. coli ArcticExpress by western blot.

Caerolysin lacks hemolytic activity against erythrocytes from different mammals

His-tagged Caerolysin was expressed in E. coli BL21 and purified from inclusion bodies over a Ni-NTA after being denatured in 8 M guanidine hydrochloride and 2 mM β- mercaptoethanol. The protein was refolded while still bound to the Ni-NTA resin in a decreasing concentration gradient of guanidine hydrochloride and was eluted with imidazole.

β-mercaptoethanol and imidazole were removed by size-exclusion chromatography followed by dialysis, and a minor fraction of Caerolysin was recovered in a soluble form (Figure 9A).

Erythrocytes from goat, sheep, cow, horse and pig were treated with pure Caerolysin but no hemolytic activity was observed when compared to the negative control (Figure 9B).

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Figure 9. CC1G_10318 does not show hemolytic activity against mammalian erythrocytes. A minor amount of His-tagged (N-terminus) CC1G_10318 was purified from inclusion bodies in a Ni-NTA column (A). In column folding was carried out with a guanidine hydrochloride gradient of decreasing concentration followed by protein elution in 200 mM imidazole. Remnants of imidazole and β-mercaptoethanol were removed by dialysis in PBS.

Fractions corresponding to the column input (Input), load-derived flow-through (FT), wash- derived flow through (Wash), imidazole elution (Elution) and post-dialysis soluble fraction (D) were mixed with Laemmli buffer, loaded on a 12% SDS-PAGE and analyzed by western blot with mouse anti-His. Pure His-tagged CC1G_10318 did not show hemolytic activity in mammal erythrocytes from different species (B).

Discussion

Serine protease inhibitors are part of the innate immune response in plants against predators

[24]. POIA1 structurally resembles a propeptide present in the serine protease Subtilisin [20].

C. cinerea encodes an ortholog of POIA1 (CC1G_06934/CCIA1) that was shown to inhibit

Subtilisin Carlsberg and truncates development in C. elegans at L4 stage. Interestingly, C. elegans encodes the Subtilisin-like peptidase Bli-4 that is essential for early development and adult morphology [35]. A single recessive mutation in Bli-4, e937, lead to a blistered phenotype in adults where the cuticle separates and the resulting space fills with liquid [36].

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Similarly, other mutated alleles of this locus cause early lethality in C. elegans larvae [37].

Bli-4 has orthologs in order nematodes, such as Necator americanus, Haemonchus contortus, Brugia malayi and Pristionchus pacificus; and has been shown to be expressed in the adult C. elegans intestine [38]. These sequence homology and tissue specificity of gene expression make Bli-4 an interesting target candidate for CCIA1 toxicity in adult worms; nonetheless, additional evidence supporting this hypothesis is needed. Loss of function mutations in the locus encoding Bli-4 will result on a phenocopy of the toxicity caused by

CCIA1 and would not provide any additional information about resistance or toxicity mechanisms. In vitro characterization of the CCIA1-Bli-4 interaction is necessary to establish a functional link between the C. cinerea inhibitor and the C. elegans peptidase.

Ricin B fold domain-containing proteins (also called β-trefoil domain-containing proteins) like

CC1G_05299 and CC1G_10603, share a common evolutionary origin [39]. A functionally diverse group of proteins, such as cytokines [40], growth factors [41], protease inhibitors [2] and lectins [6], contains this domain demonstrating that it can play multiple biological roles.

This functional plasticity is based on the large sequence variability present in the loops connecting the characteristic β-strands in the domain. β-trefoil protease inhibitors such as

Coprinopsis Cospin [2], bind to protease active sites with one or several of these loops, blocking the access of the peptidase to its substrate. Sabotic and collaborators showed that

CC1G_05299 is an inhibitor of endogenous aspartate peptidases (personal communication) and might share similar loop-mediated blockage of the catalytic site as shown for the serine protease inhibition mechanism displayed by Cospin [2].

Expression and purification of the putative pore-forming protein Caerolysin and the secreted

Ricin-B fold domain-containing protein CC1G_10603 proved to be technically demanding. In order to obtain pure recombinant proteins, different expression systems and purifications strategies were tried with modest success. Pore-forming toxins generally undergo significant conformational changes during oligomerization and pore formation [31, 42], exposing highly

134

Chapter 4 Characterization of putative defense proteins of C. cinerea hydrophobic residues that shift the protein from a soluble to an insoluble (membrane soluble) isoform. As for membrane proteins [43], these hydrophobic regions represent a challenge during folding in heterologous expression systems.

In conclusion, protease inhibitors and pore-forming toxins are interesting defense protein candidates in fungi, some of them showing entomotoxic or nematotoxic activity. The abundance of hydrophobic residues in pore-forming proteins makes their recombinant production and biochemical characterization challenging.

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18(5):581-586.

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes

Chapter 5

Toxicity spectrum analysis of fungal defense lectins in Rhabditid

and Diplogasterid nematodes

David Fernando Plaza1, Markus Aebi1, Markus Künzler1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Switzerland.

Contributions:

Toxicity assays

Intestinal staining

Lectin blots

Lectin-mediated affinity chromatography

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes

Toxicity spectrum analysis of fungal defense lectins in Rhabditid

and Diplogasterid nematodes

David Fernando Plaza1, Markus Aebi1, Markus Künzler1

1 Institute of Microbiology, Department of Biology, ETH Zürich, Zürich, Switzerland.

Abstract

With the exception of the model worm C. elegans and a few parasitic species causing human disease or agricultural losses, Nematoda represents a largely unexplored phylum.

Nematodes occupy different ecological niches with some species preying on fungi.

Accordingly, fungi have evolved an inducible innate defense system where nematotoxic lectins are produced in response to predation. In order to assess the specificity of these lectins with regard to nematode species, we fed six different strains and species of bacterivorous nematodes belonging to the orders Rhabditida and Diplogasterida (C. elegans

N2, C. elegans PMK1, C. briggsae AF16, C. sp. 11 JU1373, Distolabrellus veechi,

Halicephalobus gingivalis (deletrix) and Pristionchus pacificus) with bacteria expressing the fungal defense lectins CGL2, CGL3, CCL2, AAL, CML1, XCL, TAP1, LbTec-2, MOA and

CNL. We found that lectins targeting glycan epitopes formed by Gal linked to GalNAc, such as XCL, TAP1 and MOA were broadly toxic to nematodes. In contrast, narrow-spectrum toxicity was observed for the lectins CGL2, CCL2, AAL, CML1 and LbTec-2 that targeted only some of the species tested. CGL2 was shown to be toxic to C. elegans but innocuous to the closely related species C. briggsae and C. sp. 11. Staining of C. elegans and C. briggsae

L1 larvae with TAMRA-labelled CGL2 showed that this lectin does not bind intestinal glycans in C. briggsae. The resistance observed in the diplogasterid P. pacificus to the fucose binding lectins CCL2 and AAL was further studied by lectin blot and lectin-mediated affinity chromatography on nematode protein extracts. Glycoproteins bearing fucosylated epitopes were detected in both C. elegans and P. pacificus, suggesting that the expression of these epitopes may be organ-specific and resistance to lectins can develop by altering the

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes accessibility of some glycans to lectin binding. In conclusion, this study shows that glycan diversity in nematodes involves not only the glycan structures that can be synthesized by the enzymatic machinery encoded in their genomes, but also by the tissue-specific expression of these glycans.

Keywords: Organ-specific glycoproteome, species-specific glycoproteome, rhabditid nematodes, diplogasterid nematodes, fungal defense lectins, glycan specificity, predation.

Introduction

Little is known about the glycan repertoire of different nematode species. The most extensively studied nematode is the free-living rhabditid C. elegans. N-glycans in C. elegans mainly comprise truncated paucimannose structures [1], displaying core fucose and galactose motifs. Core 1 O-glycans consist of alpha-linked GalNAc that is attached to serine or threonine residues and further decorated with Gal, Glc or GlcA. Even though the glycomes of other nematodes have not been studied in such detail, a look at some of the genomes available, such as the genome of the necromenic diplogasterid P. pacificus, shows a significant expansion in the number of loci encoding putative glycosyltransferases in comparison to C. elegans [2].

Species-specific glycan epitopes for parasitic trematodes and nematodes of medical importance have been described [3]. For example, terminal β3-linked tyvelose is characteristic of N-glycans synthesized by the trichinosis-causing nematode Trichinella spiralis. Antibodies produced by infected patients recognizing tyvelose are protective and can be used as diagnostic tools [4, 5]. Whereas 2-O-methylated fucose is a constituent of C. elegans O-glycans [6], methylated galactose has only been observed in O-glycans from the zoonotic genus Toxocara [7]. Core β2-linked xylose, a characteristic of plant N-glycans, has been found in the trematode Schistosoma mansoni [8] but not in nematodes so far. Similarly,

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes

S. mansoni produces O-linked polymers of β3-linked GlcA and GalNAc connected to each other by β6 linkages which are not observed in other parasitic nematodes [9]. These examples reflect only a fraction of the estimated glycan diversity of nematodes or helminths in general. It is likely that the observed diversity is the result of an arms race between glycan-targeting defense proteins of the host or prey organisms and the glycans produced by the helminths.

Fungi are prey to different animals including nematodes [10]. As defense against these predators, fungi protect their reproductive organs with a battery of nematotoxic lectins showing a broad range of carbohydrate specificities [11, 12]. Some of these lectins are induced upon nematode predation suggesting that they are part of a complex innate defense system that protects fungi from foraging [11, 13]. Nematotoxicity of fungal lectins has so far only been demonstrated against the bacterivorous nematode C. elegans by feeding them E. coli cells expressing the recombinant lectins [11, 13-15]. Accordingly, the range of nematotoxicity of these fungal lectins is not known. A certain specificity can be anticipated as these lectins face a considerable diversity of nematode glycans based on some of the reports above and the huge diversity of the phylum Nematoda which comprises more than

25000 described species [16]. A large glycome diversity in nematodes can also be inferred from the non-lethality of several deletion mutations in glycosylation-related genes [13, 17], suggesting that a big proportion of these genes are dispensable under laboratory conditions and potentially implicated in ecological interactions with other species.

In this study, we show that different species of nematodes belonging to the orders Rhabditida and Diplogasterida display different susceptibility to an array of fungal defense lectins with different carbohydrate specificities. In addition, we show that in some cases, resistance is the consequence of a lack of the enzymes necessary to synthesize glycans recognized by the toxin. In other cases, we observed that lectins recognize targets in nematode-derived whole

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes glycoprotein extracts but no toxicity is evident, suggesting that the glycans recognized by these lectins are not accessible in the nematode tissues exposed to the toxin during feeding.

Methods

Nematode strains and cultivation conditions

Caenorhabditis elegans N2, C. elegans PMK1, C. briggsae AF16 and Caenorhabditis sp. 11

JU1373 (C. tropicalis) were purchased from the Caenorhabditis Genetics Center (CGC) at the University of Minnesota. P. pacificus PS312, Halicephalobus gingivalis and Distolabrellus veechi (environmental isolate) were kindly provided by Prof. Iain Wilson at BOKU University

(Vienna, Austria), Dr. Pamela Fonderie at Ghent University (Belgium) and Dr. Luis Lugones at Utrecht University (The Netherlands), respectively. Worms were grown on NGM plates (51 mM NaCl, 2.5 g/L bacteriological peptone, 13 mM cholesterol and 1.7% agar) pre-seeded with E. coli OP50 at 20°C. H. gingivalis was grown at 37°C.

Lectin cloning, expression, purification and labeling

Genes encoding untagged Coprinopsis galectin 2 (CGL2) [13], CGL3 [18], Coprinopsis cinerea lectin 2 (CCL2) [14], Aleuria aurantia lectin (AAL), Coprinopsis mucin-binding lectin

(CML1) (Bleuler S, personal communication), Xerocomus chrysenteron lectin (XCL) [11, 19,

20], Sordaria macrospora transcript associated with perithecial development (TAP1) [11],

Laccaria bicolor tectonin 2 (LbTec-2) [21], Marasmius oreades agglutinin (MOA) [15] and

Clitocybe nebularis lectin (CNL) [22] were cloned in pET expression vectors and transformed into E. coli BL21 as previously described. In addition, His-tagged versions of AAL and CCL2 were also cloned in pET vectors and transformed into E. coli BL21. Expression of His-tagged

AAL or CCL2 in bacterial transformants pregrown in LB broth (OD600: 0.5) was induced with 1 mM Isopropyl β-D-1-thiogalactopyranoside (IPTG) for 18 h at 24°C. Cells were harvested and suspended in either PBS (AAL purification) or pH 6.0 CCL2 buffer (50 mM Na2HPO4,

150mM NaCl) supplemented with 1 mM phenylmethanesulfonylfluoride (PMSF). Bacteria were lysed in a French press and centrifuged at 26000 xg to recover soluble proteins in the 147

Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes supernatant which were then bound to 1 mL Protino Ni-NTA agarose (MACHEREY-NAGEL) at 4°C (AAL) or 24°C (CCL2) under constant mixing. After washing, proteins were eluted with

200 mM imidazole, desalted in PD-10 columns (GE Healthcare) and their concentration measured by NanoDrop. CGL2 was purified in PBS on a 1 mL lactosyl-sepharose column and eluted with 200 mM lactose. Thereafter, CGL2 was dialyzed in PBS in a 10000 MWCO

Slide-A-Lyzer Dialysis Cassette (Thermo Scientific) and its concentration measured with a

NanoDrop. In order to assess the intestinal binding of CGL2 in CGL2-susceptible or resistant nematodes, CGL2 and BSA were labeled with red fluorescent Tetramethylrhodamine

(TAMRA) (Invitrogen) according to the manufacturer instructions. In brief, 10 mg CGL2 or

BSA were dissolved in 1 mL 100 mM sodium bicarbonate buffer pH 8.3 and mixed with 100

µL 10 mg/mL TAMRA dissolved in dimethyl sulfoxide. The solutions were incubated for 1 h at

24°C with continuous stirring and desalted in PD-10 columns (GE Healthcare) to remove the un-conjugated dye. Labeled protein was flash frozen in liquid nitrogen and stored at -80°C for later use.

Liquid culture of C. elegans and P. pacificus for protein extraction

C. elegans PMK1 and P. pacificus PS312 liquid cultures were carried out as previously described [23] with minor modifications: E. coli Na22 was grown for 18 h at 37°C in 2 L broth containing 12 mg/mL bacto tryptone, 24 mg/mL yeast extract, 0.4% glycerol and 50 mM

KH2PO4, and harvested by centrifugation for 5 min at 8000 xg. The bacterial pellet was flash- frozen and stored at -20°C for later use. C. elegans PMK1 and P. pacificus PS312 were grown on NGM plates pre-seeded with E. coli OP50 and harvested in sterile S-basal solution

(100 mM NaCl and 50 mM KH2PO4). Thereafter, worms were transferred to a 500 mL baffled culture flask containing 100 mL S-basal, 3 mM MgSO4, 3 mM CaCl2, 1 mL KCitrate pH 6 (1 M

Citrate and 3 M KOH), 25 µM FeSO4, 55 µM Na2EDTA, 10 µM MnCl2, 10 µM ZnSO4, 960 nM

CuSO4, 100 units penicillin, 100 µg streptomycin and 5 mL concentrated E. coli Na22; and grown for 5 days at 20°C under constant shaking (180 rpm). Worms were harvested by sucrose floatation as previously described [24]. Cultures were centrifuged for 2 min at 1200

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes xg and suspended in 22 mL 100 mM cold NaCl. 22 mL 60% sucrose were added to the worm suspension and mixed before being centrifuged for 5 min at 3800 xg. The upper most layer containing the worms was collected and transferred to a 50 mL conical bottom tube and washed three times with cold 100 mM NaCl. Worms were flash frozen and stored at -80°C.

Nematode toxicity assays

To evaluate the differential susceptibility of different worm species to fungal defense lectins, toxicity assays were performed as described earlier with small modifications [25]. NGM plates (50 mM NaCl, 2.5 g/L bacteriological peptone, 13 mM cholesterol and 1.7% agar) pre- seeded with E. coli OP50 were used to grow worms at 20-24°C until most of the bacteria was depleted and a major fraction of the worms corresponded to gravid hermaphrodites (H. gingivalis growth and toxicity assays were carried out at 37°C). To purify eggs, gravid hermaphrodites were treated with a solution containing 0.5 N NaOH and 1% NaClO for ≈ 10 min in 15 mL conical bottom tubes. Eggs were washed twice with 8 mL distilled deionized water (ddH2O) and suspended in 300-500 µL ddH2O before being transferred to a 1.5% agar plate for overnight hatching. Subsequently, L1 larvae were collected in sterile-filtered PBS, counted and adjusted to a density corresponding to 1500 worms/mL. Toxicity assays were carried out in 96-well plates using four replicates per treatment. Every well contained 20-30 worms and 180 µL lectin-expressing E. coli BL21 (initially grown to OD600: 0.5 in LB broth and induced with 1 mM IPTG for 18 h at 24°C before being adjusted with PBS to OD600: 2 and used). As a negative control, worms were fed in four replicates with pre-induced E. coli BL21 containing empty pET vector. Worms were grown for 48 h with the exception of D. veechi and P. pacificus which were grown for 72 h due to their longer life cycle. Lectin expression was confirmed by SDS-PAGE on the whole protein extracts derived from the bacterial strains used.

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Lectin-blot on C. elegans and P. pacificus whole protein extracts

To assess the presence of P. pacificus or C. elegans glycoproteins targeted by CCL2 and

AAL, whole protein extracts were prepared from frozen worms grown in liquid culture and frozen as described above. 400 mg worms were suspended in 400 µL PBS supplemented with 1X Complete Protease Inhibitor Cocktail (Roche) and 1% NP-40 detergent. Worms were homogenized in the presence of 250 mg 0.5 mm glass beads in three FastPrep FP120 steps of 6 m/s, cooling the samples on ice between steps. The extracts were centrifuged for 15 min at 12000 xg and 4°C to remove debris and insoluble proteins. The supernatants, corresponding to the PBS/NP-40-soluble whole protein extracts, were transferred to a fresh microcentrifuge tube and the protein concentration was measured by BCA. 70 µg/lane whole protein extract were loaded on a 7% SDS-PAGE and run for 2 h at 120 V. Proteins were blotted into nitrocellulose using a Trans-Blot SD semi-dry transfer cell (BioRad) and stained for two hours with 5 µg/mL His-tagged AAL or CCL2 in PBS supplemented with 0.1% Tween

20 (PBS-T). Membranes were washed twice with PBS-T and blocked with 5% bovine serum albumin (BSA). Monoclonal mouse anti-Tetra His (IgG1, Qiagen) and HRP-conjugated polyclonal goat anti-mouse IgG (LabForce AG) were used as primary and secondary antibodies, respectively, in 1:2000 dilutions. Chemiluminescence was detected by exposing an X-ray film for 2 min to the membrane treated with ECL detection mix (GE Healthcare).

CCL2-affinity chromatography of whole protein extracts from C. elegans and P. pacificus

150 µL Protino Ni-NTA agarose were equilibrated with 500 µL PBS containing 1X cOmplete

Protease Inhibitor Cocktail (Roche) and 1% NP-40 (PBS/PIC/NP-40) in Mobicol spin columns

(MoBiTec). In order to eliminate agarose- or Ni-binding proteins, C. elegans PMK1 or P. pacificus PS312 protein extracts were incubated with PBS/PIC/NP-40-equilibrated Ni-NTA for 1 h at 4°C on a rotating wheel. Unbound proteins were recovered in a collection tube by centrifugation at 0.8 xg and 4°C and loaded on fresh Mobicol spin columns containing 150 µL

Ni-NTA pre-equilibrated with 1.5 mL PBS/PIC/NP-40 and coupled to 2.5 mg His-tagged

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes

CCL2. Spin columns were incubated for 1 h at 24°C under constant mixing (rotating wheel), spinned down at 0.8 xg. Columns were washed with 1.5 mL PBS/PIC/NP-40 and eluted by incubation with 300 µL 200 mM imidazole in PBS/PIC/NP-40 for 15 min under constant mixing, followed by 1 min centrifugation at 0.8 xg. The flow through from the loading and washing steps, as well as the imidazole-eluted protein and agarose resin from the eluted columns, were mixed in a 1:1 ratio with 4X Laemmli buffer and loaded on a 10% SDS-PAGE that was silver-stained thereafter. To control that proteins observed by silver staining were not worm-derived histidine-rich proteins which were un-specifically bound to the Ni-NTA column, the same fractions were run on a second 10% SDS-PAGE, spotted on nitrocellulose, and detected using the same anti-His antibody combination and concentration as in the lectin blot described above.

Intestinal staining with fluorescently-labelled CGL2

To study the intestinal binding of CGL2 to glycans in C. elegans and C. briggsae, nematodes were grown on NGM plates pre-seeded with B. subtilis 168. Synchronized C. elegans N2 or

C. briggsae AF16 L1 larvae (the offspring of worms grown on B. subtilis 168) were incubated for 2 h in 100 µl PBS supplemented with 500 µg/ml TAMRA-labeled CGL2 or BSA and B. subtilis (OD600: 2) at 24°C under constant shaking (500 rpm). Worms were washed three times with 1 mL PBS (1300 xg for 1 min at 24°C) to remove the excess of protein and incubated with B. subtilis (OD600: 2) in 100 µL PBS under constant shaking (500 rpm) at

24°C for 18 h in order to remove unbound protein. Worms were centrifuged at 1300 xg for 1 min, suspended in 20 µL PBS supplemented with 100 mM levamisole and mounted on microscope slides.

Results

Fungal defense lectins show a differential toxicity against different nematode species

Toxicity against C. elegans Bristol N2 has been previously demonstrated for lectins produced by C. cinerea, S. macrospora, X. chrysenteron, Clitocybe nebularis, A. aurantia, Sclerotinia

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes

sclerotiorum and M. oreades [11, 13-15]. To study

how broadly toxic these lectins are against

nematodes other than C. elegans, we fed L1 larvae

from P. pacificus, H. gingivalis, D. veechi, and C.

briggsae with E. coli BL21 expressing untagged

forms of the fungal lectins CGL2, CGL3, CCL2, AAL,

CML1, XCL, TAP1, LbTec-2, MOA and CNL,

respectively. We found that some lectins previously

described to be toxic against C. elegans such as

CGL2, CCL2, AAL and LbTec-2 were not toxic

against some other nematode species (Figure 1). In

contrast, lectins recognizing glycan epitopes

composed by Gal and GalNAc, such as XCL, TAP1

and MOA, showed broad-spectrum toxicity against all

the nematode species and strains tested suggesting

that this structures and the tissue regulation of the

enzymes that synthesize them are conserved among

nematodes (Figure 1). Similarly, the LacNAc- and

chitobiose-specific CGL3, whose structure is very

similar to CGL2 but did not show any toxicity to C.

elegans, as well as the LacdiNAc-specific CNL, were

not toxic to any of the other nematodes suggesting

that these epitopes are either not present or not

accessible in the digestive tract of these animals.

These results suggest that fungi produce both broad-

and narrow-spectrum defense lectins. The spectrum

of a specific lectin possibly depends on the

conservation of the respective target glycans among

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes nematodes.

Figure 1. Broad- and narrow-spectrum lectins as defense effectors of fungi against nematodes. L1 larvae from different strains of rhabditid and diplogasterid nematodes were fed with lectin-expressing E. coli BL21. Toxicity is indicated as '+' and green color while resistance is marked by '-' and red color. Reported glycan specificities with the corresponding reference are shown for the different lectins used. N/A: toxicity data not available.

AAL and CCL2 bind carbohydrate epitopes present in the glycoproteome of P. pacificus.

To assess the presence or absence of P. pacificus proteins decorated with carbohydrate structures targeted by the non-toxic lectins AAL and CCL2, a lectin blot was carried out on the C. elegans PMK1 and P. pacificus PS312 PBS/NP-40-soluble protein extracts. AAL and

CCL2 were shown to specifically recognize numerous glycoproteins in the extracts derived from both nematode species (Figure 2A). In accordance with these results, CCL2-affinity chromatography specifically captured glycoproteins from both nematode species; however, different bands were found to be recognized by CCL2 in the two proteomes (Figure 2B). The proteins eluted from the column were not rich in histidine (Figure 2C), suggesting that the enriched proteins from C. elegans and P. pacificus are specifically bound by CCL2.

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes

Figure 2. P. pacificus glycoproteins are recognized by the fucose-binding lectins AAL and CCL2 (A) AAL and CCL2 lectin blots on protein extracts from C. elegans PMK1 (Ce) and P. pacificus PS312 (Pp). 70 µg/lane NP-40-soluble protein extracts from Ce and Pp were spotted on nitrocellulose membranes and stained with His-tagged AAL or CCL2. Monoclonal mouse anti-His and HRP-coupled goat anti-mouse IgG were used as primary and secondary antibodies, respectively, to detect the lectins. As controls, protein extracts were overlaid with primary and secondary antibodies only (anti-His) i.e. without the lectins. (B) and (C). CCL2- affinity chromatography of NP-40 soluble protein extracts from Ce and Pp. NP-40-soluble protein extracts from Ce and Pp were depleted of agarose and Ni-binding proteins, and loaded into Ni-NTA columns preloaded with His-tagged CCL2 and washed. The flow-through from the loading (FT) and wash (Wash) steps were collected and CCL2-bound nematode 154

Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes glycoproteins were eluted in 200 mM imidazole (Eluate). In addition to the three fractions, the eluted agarose beads (Beads) were mixed with 4X Laemmli buffer and run in two 10% SDS-

PAGEs. One of the gels was silver-stained to detect proteins bound to CCL2 (B). Arrows indicate species-specific proteins captured by CCL2. The second gel (C) was spotted on a nitrocellulose membrane and stained with mouse anti-His and HRP-coupled goat anti-mouse

IgG as primary and secondary antibodies, respectively, to detect His-rich proteins in the nematode extracts. Relative concentrations of the fractions loaded are shown at the bottom of panels (B) and (C).

CGL2 does not bind to the intestinal epithelium of C. briggsae AF16

Previous studies have shown that the nematotoxicity of fungal lectins is mediated by the binding to glycan epitopes in the worm intestine [13, 14]. To test whether the CGL2- resistance observed in C. briggsae and C. sp. 11 was caused by a lack of the enzymatic machinery necessary to synthesize the CGL2 target glycan, a BLAST-search was performed on the genome of these two species for orthologous enzymes to those known in C. elegans to be responsible for the synthesis of the N-glycan target of CGL2 [13] (Figure S1).

Intriguingly, highly similar orthologous enzymes of Bre-1 (CBG21737 and

Csp11.Scaffold629.g10870.t1), Gly-13 (CBR-GLY-13 and Csp11.Scaffold629.g13419.t1),

Aman-2 (CBR-AMAN-2 and Csp11.Scaffold629.g7777.t1, the latter annotated using RNA- seq data kindly provided by Michael Paulini), Fut-8 (CBR-FUT-8 and

Csp11.Scaffold491.g2080.t1) and Galt-1 (CBR-GALT-1 and Csp11.Scaffold491.g2092.t2) were found in the genomes of C. briggsae AF16 and C. sp. 11 JU1373.

To assess the presence of the so-called Gal-Fuc epitope recognized by CGL2 in the intestinal epithelium of C. briggsae, purified CGL2 was fluorescently-labeled with TAMRA and used to feed C. elegans N2 and C. briggsae. Thereafter, the unbound protein was washed out from the intestine by feeding the worms with B. subtilis 168. Intestinal fluorescence was observed in C. elegans larvae but not in larvae of C. briggsae (Figure 3),

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes suggesting that the glycan targeted by CGL2 is not available in the intestine of C. briggsae, although the genome of this worm encodes the entire enzymatic machinery required for its biosynthesis (Figure S1).

Figure 3. The intestine of C. briggsae AF16 is not stained by the Coprinopsis galectin

CGL2. C. elegans and C. briggsae L1 larvae were fed with 500 µg/mL TAMRA-labeled CGL2 and B. subtilis 168 for 2 h. The excess of lectin was removed and the unbound CGL2 was washed out from the intestine by feeding the worms for 18 h with B. subtilis 168 only. In contrast to C. elegans, CGL2 intestinal staining was not observed in C. briggsae larvae. As a negative control, larvae were fed with 500 µg/mL TAMRA-labeled BSA. BSA staining was not observed in any of the worms. PC: Phase contrast. RF: Red fluorescence channel.

C. sp11 JU1373 may lack one enzyme required in the biosynthesis of the Gal-Fuc epitope recognized by CGL2

In contrast to the orthologs present in C. elegans and C. briggsae, the predicted amino acid sequence of Galt-1 (Csp11.Scaffold491.g2092.t2) from C. sp. 11 contains a deletion of 8 amino acid residues belonging to the only galactosyltransferase domain present in the protein (Figure S1). This deletion might affect the function or substrate specificity of Galt-1 rendering C. sp. 11 resistant to CGL2 [13].

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes

Discussion

The differential susceptibility to fungal lectins observed in a group of closely related species of nematodes suggests that there is a large glycan diversity in the phylum Nematoda that has not been explored. This diversity could be derived from alterations in glycan structure or tissue specific expression which might not affect the ecological fitness of the worms.

The susceptibility differences observed in species of the genus Caenorhabditis might reveal how some of its glycome diversity could have arisen. In contrast to C. elegans, C. briggsae

AF16 and C. sp. 11 JU1373 were resistant to the N-glycan-linked galactose-recognizing lectin CGL2. Loss-of-function mutations in enzymes responsible for glycoepitope biosynthesis could cause resistance to lectins in nematodes. For instance, C. sp. 11 resistance to CGL2 might be partially explained by a loss-of-function mutation in the galactosyltransferease Galt-1 (Figure S1). It was shown that C. elegans mutants lacking galt-1 display an altered spectrum of N-glycans and are unable to produce the Gal-Fuc epitope on N-glycan cores shown to be the target of CGL2 in this nematode [13], supporting the idea that C. sp. 11 lacks the enzymatic machinery necessary to synthesize the glycan epitope recognized by CGL2.

In contrast, a lack of intestinal staining with CGL2 in C. briggsae cannot be explained by an absence of enzymes responsible for the biosynthesis of this glycan epitope in the genome of this species. A possible explanation for this observation is that the epitope recognized by this lectin is not expressed in the intestine but other tissues in the worm. Tissue-specificity of glycan expression has been poorly studied in nematodes. A staining comparison with the plant lectins (specificities shown in parenthesis) ConA (Man-rich [26]), RCA-II (Gal or GalNAc

[27]), PHA-E (biantennary galactosylated N-glycan with bisecting GlcNAc [28]), PHA-L (β1-6

GlcNAc branching structures and tetraantennary complex type oligosaccharides [29]), LCA

(core Fuc [30]), STL (GlcNAc [27]), UEA-I (α2-linked Fuc [31]), PNA (Galβ1-3GalNAc [31]),

SBA (Galβ1-4GlcNAc [32]) and LPA (Neu5Ac [33]) was performed in the rhabditid

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Chapter 5 Toxicity spectrum of fungal defense lectins in nematodes nematodes C. elegans, Panagrolaimus superbus and Acrobeloides maximus [34]. Nematode inter-species differences were only observed for the organ distribution of Galβ1-3GalNAc as shown by the differential staining with PNA of neurons in C. elegans or egg yolk in P. superbus and A. maximus [34]. In mouse, tissue-specific N-glycosylation sites have been detected by filter aided sample preparation (FASP) coupled to LC-MS [35]. Furthermore, the expression of glycosylation-associated genes (glycogenes) measured by microarrays has been shown to vary significantly between different organs in mouse, where only 15% of the

470 glycogenes studied were found to be expressed in all the organs sampled (bone marrow, brain, kidney, liver, lymph nodes, lung, spleen, testes and thymus) [36]. This patchiness of gene expression involved in glycan biosynthesis in mammals has been confirmed by qRT-PCR [37] and suggests that glycan epitope availability differs from organ to organ in the same organism.

In summary, this study shows that fungi produce broad- and narrow-spectrum defense lectins against nematodes. Nematode resistance to lectin toxicity can be explained by a lack of lectin ligands in the nematode intestine as shown in C. briggsae fed with TAMRA-labelled

CGL2. In contrast, the absence of toxicity targets in C. sp. 11 might be due to conspicuous changes in the galactosyltransferase domain of Galt-1 that is involved in the synthesis of the galactosylated N-glycan epitope bound by CGL2 in C. elegans. Intriguingly, resistance to the fucose binding lectins AAL and CCL2 in the diplogasterid nematode P. pacificus is not derived from a lack of binding epitopes as it is concluded from the binding of these lectins to several glycoproteins in the glycoproteome of this worm. Presence of fucosylated epitopes is demonstrated by the CML1-derived toxicity observed in P. pacificus. Further evidence is necessary to mechanistically describe the toxicity determinants of fungal lectins, however, inter- and intra-species diversity in glycan structure and glycogene expression in nematodes might partially explain why some fungivorous nematodes can feed on lectin-expressing fungi while others cannot.

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Acknowledgements

We are grateful to Prof. Iain Wilson at BOKU University in Vienna, Dr. Pamela Fonderie at

Ghent University and Dr. Luis Lugones at Utrecht University for kindly providing P. pacificus,

H. gingivalis and D. veechi. We also thank Andrea Ochsner and Philipp Moosmann for performing preliminary experiments during this study. This project was supported by the

Swiss National Science Foundation Grant 31003A_130671.

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Supplementary information

Figure S1. Genes known to be involved in the synthesis of the N-glycan epitope recognized by CGL2 in C. elegans are highly conserved in C. briggsae and C. sp. 11 Multiple sequence alignments were carried out in the CLC Genomics Workbench suite. Synthesis diagram was adapted from Butschi A et al, 2010.

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Bre-1

Gly-13

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Aman-2

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Fut-8

Galt-1

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Chapter 6 Conclusions

Chapter 6

Conclusions and outlook

169

Chapter 6 Conclusions

Conclusions

The current understanding of the fungal immune system is comparable to the one we had of our own by the end of the 19th century. By then, we knew there were effectors, now known as antibodies [1] and complement [2], that were present in the human body fluids and that had antimicrobial activity specific to some microorganisms but not to others. We also knew that some of these effectors were induced upon infection or surgical tissue damage but nothing was really known about their precise regulation mechanisms, the cell type responsible for their production or the basis of their specificity. Similarly, in filamentous fungi we have recently discovered that cytoplasmic proteins specific to glycoepitopes not expressed by the fungal producer, belong to a non-self recognition system against predator insects and nematodes [3]. In addition, these proteins extended the defense repertoire of fungi that includes well known antibacterial secondary metabolites [4] and proteins [5]. In the course of this project I demonstrated that these lectins and antibacterial proteins are developmentally regulated in Coprinopsis cinerea in order to protect mushrooms and vegetative hyphae from predators and competitors that inhabit the environment. Moreover, I showed that defense lectins and putative antibacterial proteins of C. cinerea are specifically induced in vegetative hyphae exposed to fungivorous nematodes or bacteria, respectively, indicating that the fungal immune system can discriminate between different types of environmental signals and regulates gene expression in order to respond to these cues.

On the side of the nematode predators, the data revealed that the evolution of a glycan- targeting defense system in fungi might have led to the diversification of the glycorepertoire in worms, giving rise to interesting patterns of resistance or susceptibility in closely related species. Lectin blots and in vivo lectin staining using susceptible and resistant species of nematodes will provide conclusive evidence on the presence or not of lectin recognition glycoepitopes and their tissue-specific regulation in these species.

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Chapter 6 Conclusions

In conclusion, this project showed that the co-evolution of fungi and their nematode predators has led to the development of a complex fungal innate immune system that uses lectins, protease inhibitors and putative antibacterial peptides as defense effectors. Moreover, the expression of proteins associated to this defense system was shown to be transcriptionally regulated by developmental and environmental cues. Finally, the evolutionary arms race between fungi and their predators is the most likely driving force (selective pressure) behind the large diversity existing of fungal lectin specificities and nematode glycorepertoires in nature.

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