Cell Wall Biosynthesis in the Pathogenic Oomycete Saprolegnia Parasitica

Cell wall biosynthesis in the pathogenic oomycete Saprolegnia parasitica

Elzbieta Rzeszutek

Doctoral Thesis in Biotechnology

School of Engineering Sciences in Chemistry, Biotechnology and Health Royal Institute of Technology Stockholm, Sweden 2019

ISBN: 978-91-7873-370-5 TRITA-CBH-FOU-2019:61

KTH School of Engineering Sciences in Chemistry, Biotechnology and Health Royal Institute of Technology AlbaNova University Center 106 91 Stockholm Sweden

Printed by Universitetsservice US-AB, Stockholm 2019 Abstract The oomycete Saprolegnia parasitica is a fungus-like microorganism responsible for the fish disease saprolegniosis, which leads to important economic losses in aquaculture. Currently, there is no efficient method to control the infection and therefore methods for disease management are urgently needed. One of the promising approaches to tackle the pathogen is the inhibition of cell wall biosynthesis, specifically the enzymes involved in carbohydrate biosynthesis. The cell wall of S. parasitica consists mainly of cellulose, β-1,3 and β-1,6-glucans, whereas chitin is present in minute amounts only. The available genome sequence allowed the identification of six putative chitin (Chs) and cellulose (CesA) synthase genes. The main objective of this work was to characterize CHSs and CesAs from S. parasitica and test the effect of cell wall related inhibitors on pathogen growth. The tested inhibitors included nikkomycin Z, a competitive inhibitor of CHS as well as inhibitors of cellulose biosynthesis, namely flupoxam, CGA325'615 and compound I (CI). All drugs strongly reduced the growth of S. parasitica and inhibited the in vitro formation of chitin or cellulose, demonstrated by the use of a radiometric assay. The chemicals also affected the expression of some of the corresponding Chs and CesA genes. One of the CHSs, namely SpCHS5, was successfully expressed in yeast and purified to homogeneity as a full length protein. The recombinant enzyme was biochemically characterized and demonstrated to form chitin crystallites in vitro. Moreover, our data indicate that SpCHS5 most likely occurs as a homodimer which can further assemble into larger multi- subunit complexes. Point mutations of conserved amino acids allowed us to identify the essential residues for activity and processivity of the enzyme. In addition to the cell wall related inhibitors, a biosurfactant naturally produced by Pseudomonas species, massetolide A, was tested, showing strong inhibition of S. parasitica growth. Altogether, our data provide key information on the fundamental mechanisms of chitin and cellulose biosynthesis in oomycetes and the biochemical properties of the enzymes involved. They also demonstrate that the enzymes involved in cell wall biosynthesis represent promising targets for anti-oomycete drugs, even when the corresponding polysaccharides, such as chitin, occur in small amounts in the cell wall. Keywords: Cellulose biosynthesis; chitin biosynthesis; cellulose synthase genes; chitin synthase genes; oomycetes; Saprolegnia parasitica

Sammanfattning Oomyceten Saprolegnia parasitica är en svampliknande mikroorganism som orsakar fisksjukdomen saprolegniosis, vilken leder till stora ekonomiska förluster inom vattenbruk. För närvarande finns det ingen effektiv metod för att kontrollera infektionen och därför är behovet av metoder för att begränsa sjukdom stort. Ett lovande tillvägagångssätt för att angripa patogenen är genom att hämma biosyntes av cellväggen, särskilt de enzymer som är involverade i kolhydratbiosyntes. Cellväggen hos S. parasitica består huvudsakligen av cellulosa, ß-1,3 och ß-1,6- glukaner, samt små mängder kitin. Den tillgängliga genomsekvensen möjliggjorde identifiering av sex förmodade kitin (Chs) och cellulosa (CesA) -syntasgener. Huvudsyftet med denna avhandling var att karakterisera CHS och CesA från S. parasitica och testa effekten av cellväggsrelaterade inhibitorer på patogenens tillväxt. De testade inhibitorerna inkluderade nikcomycin Z, en kompetitiv inhibitor av CHS, samt flera inhibitorer av cellulosabiosyntes, nämligen flupoxam, CGA325'615 och compund I (Cl). Alla inhibitorer reducerade kraftigt tillväxten av S. parasitica och genom användning av en radiometrisk analysmetod visades att in vitro-bildningen av kitin och cellulosa inhiberades. Kemikalierna påverkade också uttrycket av några av Chs- och CesA-generna. En av CHS, nämligen SpCHS5, uttrycktes framgångsrikt i jäst och renades till homogenitet som ett fullängdsprotein. Det rekombinanta enzymet karakteriserades biokemiskt och visade sig bilda kitinkristalliter in vitro. Dessutom indikerar våra data att SpCHS5 troligen är en homodimer som kan bilda större komplex bestående av flera subenheter. Punktmutationer av konserverade aminosyror tillät oss att identifiera de aminosyror som är väsentliga för enzymets aktivitet och processivitet. Förutom de cellväggsrelaterade inhibitorerna testades även biosurfaktanten massetolid A som produceras naturligt av Pseudomonas- arter. Denna visade sig ha en starkt inhiberande verkan på S. parasitica- tillväxt. Sammantaget bidrar våra data med viktig information rörande de grundläggande mekanismerna för biosyntes av kitin och cellulosa i oomyceter och de biokemiska egenskaperna av de involverade enzymerna. Resultaten visar också att enzymerna som är involverade i biosyntes av cellväggen är lovande mål för bekämpningsmedel mot oomyceter även då motsvarande polysackarider, såsom kitin, förekommer i små mängder i cellväggen. Nyckelord: Cellulosabiosyntes; kitinbiosyntes; cellulosasyntasgener; kitin- syntasgener; Oomyceter; Saprolegnia parasitica ii

List of publications

Publication I Jiang, R. H., de Bruijn, I., Haas, B. J., Belmonte, R., Löbach, L., Christie, J., van den Ackerveken, G., Bottin, A., Bulone, V., Díaz-Moreno, S. M., Dumas, B., Fan, L., Gaulin, E., Govers, F., Grenville-Briggs, L. J., Horner, N. R., Levin, J. Z., Mammella, M., Meijer, H. J. G., Morris, P., Nusbaum, C., Oome, S., Phillips, A. J., van Rooyen, D., Rzeszutek, E., Saraiva, M., Secombes, C. J., Seidl, M. F., Snel, B., Stassen, J. H. M., Sykes,S., Tripathy, S., van den Berg, H., Vega-Arreguin, J. C., Wawra, S., Young, S. K., Zeng, Q.,Dieguez-Uribeondo, J., Russ, C., Tyler, B. M., and Pieter van West (2013). Distinctive expansion of potential virulence genes in the genome of the oomycete fish pathogen Saprolegnia parasitica. PLoS Genetics, 9(6), e1003272.

Publication II Rzeszutek, E., Díaz-Moreno, S. M., and Bulone, V. (2019). Identification and characterization of the chitin synthase genes from the fish pathogen Saprolegnia parasitica. Accepted, Frontiers in Microbiology.

Publication III Rzeszutek*, E., Díaz-Moreno*, S. M., Ampomah, O. Y., Inman, A., Sirvastava, V., Zhou, Q., and Bulone, V. (2019). Novel insights into chitin biosynthesis through heterologous expression and biochemical characterization of chitin synthase 5 from the pathogenic oomycete Saprolegnia parasitica. Manuscript.

Publication IV Rzeszutek, E., Díaz-Moreno, S. M., Klinter, S., and Bulone, V. (2019). Analysis of the cellulose synthase genes in the oomycete fish pathogen Saprolegnia parasitica and effect of cellulose biosynthesis inhibitors on enzyme activity and microbial growth. Manuscript.

Publication V Liu, Y., Rzeszutek, E., van der Voort, M., Wu, C.-H., Thoen, E., Skaar, I., Bulone, V., Dorrestein, P. C., Raaijmakers, J. M., and De Bruijn, I. (2015). Diversity of aquatic Pseudomonas species and their activity against the fish pathogenic oomycete Saprolegnia. PloS One, 10(8), e0136241.

*Both authors contributed equally to the work

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Author’s contribution to the publications

Publication I Elzbieta Rzeszutek identified putative Chs genes in the genome of S. parasitica.

Publication II Elzbieta Rzeszutek performed the sequence and qPCR analyses of CHSs from S. parasitica, and contributed to the Southern blot experiments. She performed in vitro inhibitory assays, microscope observations, biochemical assays and characterization of the in vitro product. Elzbieta prepared most of the figures and wrote the manuscript with input from all authors.

Publication III Elzbieta Rzeszutek performed the expression, purification and biochemical characterization of SpCHS5. She also characterized the in vitro product by enzymatic treatment and glycosidic linkage analysis, and performed cell wall analysis of the yeast S. cerevisiae. Elzbieta prepared the figures and wrote the manuscript together with contributions from the other co-authors.

Publication IV Elzbieta Rzeszutek identified putative CesAs genes in S. parasitica and contributed to the qPCR experiment. She performed in vitro inhibitory assays, microscope observations, biochemical assays and in vitro product characterization. Elzbieta also contributed to the preparation of the figures and wrote the manuscript with contributions from all authors.

Publication V Elzbieta Rzeszutek contributed to the in vitro inhibitory assays and data analysis.

List of Abbreviations

Bcs Bacterial cellulose synthase CAZy Carbohydrate-active enzymes CesA Cellulose synthase CDB Carbohydrate-binding domain CHS Chitin synthase CI Compound I CLP Cyclic lipopeptides COS Chito-oligosaccharides CWP Cell wall protein DCB 2,6-dichlorobenzonitrile DMS Dimethyl suberimidate GlcNAc N-acetylglucosamine GPI Glycosylphosphatidylinositol GS Glucan synthase GT Glycosyltransferase MIT Microtubule interacting and trafficking domain NZ Nikkomycin Z PH Plekstrin homology domain SuSy Sucrose synthase TC Terminal complex TM Transmembrane domain UDP Uridine diphosphate WAE Wall-associated enzymes

Contents Introduction to oomycetes ...... 1 Saprolegnia parasitica ...... 3 Life cycle of Saprolegnia parasitica ...... 4 Control measures of Saprolegnia parasitica ...... 5 The cell wall ...... 6 Oomycete cell walls ...... 7 Fungal cell walls ...... 9 Chitin ...... 12 Cellulose ...... 13 β-1,3-Glucans ...... 15 Polysaccharide biosynthesis ...... 16 Glycosyltransferases ...... 16 Catalytic mechanisms ...... 17 Chitin synthases ...... 19 Cellulose synthases ...... 25 β-1,3-Glucan synthase ...... 30 Cell wall biosynthetic enzymes as potential targets for anti-oomycete drugs ...... 31 Cell wall related inhibitors ...... 31 Biosurfactants as new promising inhibitors for the control of saprolegniosis ...... 36 Aims of the thesis ...... 38 Results and discussion ...... 39 Paper I: Analysis of the genome of S. parasitica ...... 39 Paper II: Characterization of chitin synthases from Saprolegnia parasitica ...... 40 vi

Paper III: Functional and biochemical characterization of SpCHS5 ...... 43 Paper IV: Characterization of the cellulose synthase genes in S. parasitica ...... 46 Paper V: Activity of aquatic Pseudomonas species against Saprolegnia species ...... 48 Conclusions and research perspectives ...... 50 Acknowledgements ...... 52 References ...... 53

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Introduction to oomycetes

The oomycetes constitute a large group of fungus-like eukaryotic microorganisms. Both fungi and oomycetes have saprotrophic and/or necrotrophic life styles, and show similar mycelial growth patterns. For a long time, oomycetes were considered as true fungi (Phillips et al., 2008; Latijnhouwers et al., 2003). However, despite their morphological and life style similarities, molecular and biochemical analyses separated oomycetes from fungi and classified the oomycetes into the Stramenopiles group, together with diatoms and brown algae (Figure 1) (Beakes et al., 2012).

Fig. 1. Phylogenetic tree representing the evolutionary relationship between major eukaryotic groups (adapted from Latijnhouwers et al., 2003).

There are several important biological features that distinguish oomycetes from fungi. One of them is their cell wall composition. Oomycete cell walls consist mainly of β-1,3-glucans, β-1,6-glucans and cellulose rather than chitin, which is one of the essential constituents of fungal cell walls. Additionally, oomycete hyphae are generally aseptated (without cross walls) and contain diploid nuclei in the vegetative state.

Fungi on the other hand are typically haploid, apart from diploid zygotes and, unlike oomycetes, their hyphae generally consist of multiple cells separated by cross-walls (septae), with each cell containing one or more nuclei. During sexual reproduction, the oomycetes develop diploid oospores, which are zygotes formed by fertilization of oospheres. Moreover, they produce motile zoospores with two types of flagella, in contrast to fungi which rarely produce motile spores. In addition, fungal spores are monoflagellated (Rossman and Palm, 2006).

Table 1. Major differences between oomycetes and true fungi.

Feature Oomyctes Fungi Cell wall Cellulose, β-1,3 and β-1,6 Usually chitin, β-1,3 and β- composition glucans, small amount of 1,6 glucans; chitin in some species no cellulose Hyphal Non-septate and Septate hyphae, with one architecture multinucleate hyphae or more nuclei per compartment Nuclear state of Diploid Haploid or dikaryotic vegetative mycelium Sexual Oospores, formed by Oospores are not reproduction fertilization of oospheres by produced; sexual nuclei from antheridia reproduction results in zygospores, ascospores and basidiospores Flagella type on Biflagellated: one whiplash, If flagellated, usually one zoospores directed posteriorly; the type: whiplash directed other is fibrous and ciliated posteriorly and directed anteriorly Mitochondria Tubular cristae Flattened cristae (Adapted from Rossman and Palm, 2006; Judelson and Blanco, 2005).

Oomycetes contain many important pathogenic species responsible for devastating diseases in plants and animals (Phillips et al., 2008; Latijnhouwers et al., 2003). A well known example is Phytophthora infestans, one of the most studied plant pathogens, which is the cause of

2 potato late blight and agricultural losses of about 6 billion euros per year globally (Haas et al., 2009). Other examples are aquatic oomycetes like Aphanomyces astaci, which causes crayfish plague, and Saprolegnia species like Saprolegnia parasitica and Saprolegnia diclina, which infect fresh-water fish and eggs (Phillips et al., 2008; van West, 2006).

Saprolegnia parasitica

The oomycete S. parasitica is one of the most severe fish pathogens responsible for the infection of fresh-water fish like salmon, trout and catfish as well as fish eggs. It brings millions of euros of losses yearly in the fish farm industry and has a huge impact on natural ecosystems, decreasing the population of wild salmon (van West, 2006; van den Berg, et al., 2013). The disease caused by Saprolegnia is called saprolegniosis and it is characterized by white or grey patches of mycelium on the fish (Figure 2).

Fig. 2. Bony bream affected by Saprolegnia parasitica (from van West, 2006).

The pathogen destroys mainly epidermal layers, but it is also able to grow deeper into the underlying tissues and, if not treated, it subsequently leads to the death of the fish (van West, 2006). S. parasitica acts as an opportunistic pathogen (secondary invader). It attacks fish with weakened resistance, e.g. wounded or already infected by another

3 pathogen. Although healthy fish are rather resistant to saprolegniosis, some very virulent strains of S. parasitica can cause primary infection in healthy fish (Meyer, 1991; van West, 2006).

Life cycle of Saprolegnia parasitica

The life cycle of S. parasitica consists of both asexual and sexual stages (Figure 3; Phillips et al., 2008).

Fig. 3. Life cycle of Saprolegnia parasitica including asexual and sexual phases (from Phillips et al., 2008).

In the first step of asexual reproduction, biflagellated primary zoospores are produced in the sporangium (formed at the end of the hyphae), and subsequently released into the water. The primary zoospores rapidly encyst into primary cysts, from which secondary zoospores develop. The

4 secondary zoospores have the ability to remain motile for a longer time and afterwards encyst into secondary cysts. The later form structures called “boat hooks”, which are suggested to enhance the attachment of the cysts to the fish skin (Beakes, 1983; Pickering and Willoughby, 1982). In the absence of a suitable host, the secondary cysts are able to release new zoospores and encyst again in a process called polyplanetism (Beakes, 1983; van West, 2006; Phillips et al., 2008). This process increases the probability of the pathogen to locate a host. Once a suitable host is found, secondary zoospores germinate into new mycelium and start the infection. During the sexual phase, male antheridia and female oogonia fuse together to produce oospores, which afterwards germinate and develop into hyphal cells (Phillips et al., 2008).

Control measures of Saprolegnia parasitica

S. parasitica is responsible for important economic and environmental losses and constitutes a significant problem for the aquaculture industry (van West, 2006). In the past, Saprolegnia infections in fish and fish eggs were successfully treated with an organic dye, malachite green. However, due to the accumulation of this chemical and derived metabolites in fish tissues, their carcinogenic and toxicological effects, malachite green was banned worldwide in 2002 (Plakas, 1996; Srivastava, 2004). As an alternative, formalin has been used as an efficient treatment of oomycete infections and has been shown to reduce mortality in fish infected by S. parasitica (Gieseker et al., 2006). However, the use of formalin raised safety concerns for human health. It was not only shown to have a negative impact on water quality but also to be toxic to the fish at higher concentrations. Therefore, it has been limited or even banned in several countries (Ali et al., 2015; Leal et al., 2016). One of the safer replacements for malachite green and formalin is bronopol (2-bromo-2-nitropropane-1,3-diol). This biocide has a broad range of antimicrobial activities and is used as a preservative in many types of cosmetics and pharmaceutical products. In fish farms, bronopol

5 is effectively used to protect fish and fish eggs from contracting saprolegniosis, but it is less active against an already established infection. Thus, bronopol is not an ideal solution. In addition, its use and disposal in the aquaculture industry tend to be expensive (Picón- Camacho et al., 2012; Pottinger and Day, 1999). Additionally to the chemicals mentioned above, there are other potential solutions, but their efficacy is limited. These include hydrogen peroxide, sodium chloride or ozone treatment, which were all reported to reduce oomycete infections, eggs mortality and improve hatching rates of fish eggs. Nevertheless, compared to formalin or bronopol, they appear to be less effective (Ali et al., 2015; Forneris et al., 2003; Schreier et al., 1996). One of the interesting alternatives against saprolegniosis is the development of vaccines. Even though no vaccine against S. parasitica is currently available, a putative serine protease (SpSsp1) was identified as a promising protein for future vaccination. This protein was shown to be highly expressed in S. parasitica’s mycelium and antibody response was detected in several non-infected fish (Minor et al., 2014). Despite these possible different strategies and approaches, there is currently no ideal solution for the control of Saprolegnia diseases. Therefore alternative methods to suppress the disease have to be established. One of the promising approaches is the inhibition of oomycete cell wall biosynthesis as a target for disease control.

The cell wall

The cell wall is an external barrier surrounding the plasma membrane of plant, bacterial, fungal and oomycete cells, but it is absent in animal cells. It consists mainly of polysaccharides and proteins. However, the composition and structure vary between different organisms and depend on the cell type and development stages. In plants, the main component of the cell wall is cellulose, in bacteria peptidoglycan, and in fungi β-1,3 glucans. The cell wall is involved in many essential functions. It maintains the shape of the cell and confers mechanical protection to changes in osmotic pressure and other environmental stresses. It also provides protection against pathogenic microorganisms. In spite of its apparent

6 rigidity, the cell wall is a very dynamic structure, since it has to adapt to many modifications and changes which continuously occur during cell growth and division. In addition to its structural function, the cell wall allows interactions with the environment. It contains molecules involved in many biological processes like signalling, uptake of nutrients and, in pathogenic fungi and oomycetes, in host recognition, adhesion and colonization (Klis et al., 2006; Bowman and Free, 2006; Free, 2013). The formation and remodeling of the cell wall require numerous enzymes, including polysaccharide synthases, transglycosylases and hydrolases (Bernard and Latge, 2001). Disruption of the cell wall structure affects the growth and morphology of cells, quite often leading to cell lysis and death. In the case of the pathogenic oomycete Phytophthora infestans, interruption of cellulose synthesis results in unsuccessful infection of potato leaves (Grenville-Briggs et al., 2008). Therefore, the cell wall is an attractive target for new anti-microbial drugs.

Oomycete cell walls

Oomycete cell walls consist mainly of cellulose, β-1,3 and β-1,6 glucans (Bartnicki Garcia 1968; Mélida et al., 2013). Unlike fungi, which are devoid of cellulose, oomycetes contain cellulose as a main cell wall component, representing at least 30% of the total cell wall carbohydrates. In S. parasitica cell wall, the total amount of cellulose is about 50% (Mélida et al., 2013). Cellulose is a linear polymer of β-1,4-linked glucosyl residues and is considered to have a similar structural function as chitin in fungi. Ultrastructural studies of the Achlya bisexualis cell wall showed that cellulose is distributed along the hyphae, except for the apical region, which contains mainly β-1,3-glucans (Shapiro and Mullins, 2002(a); Shapiro and Mullins, 2002(b)). In addition, cellulose in the S. monoica cell wall exhibits a microfibrillar structure and was characterized by X-ray diffraction as a poorly crystalline form corresponding to cellulose IV (Bulone et al., 1992). As opposed to fungi, chitin is a minor component of the cell wall of oomycetes and has been detected only in a few species belonging to the Leptomitales (Lin and Aronson, 1970; Aronson and Lin, 1978) and Saprolegniales orders (Campos-Takaki et al., 1982; Bulone et al., 1992; Mélida et al., 2013). Despite the small amount of chitin present (< 2% of

7 the cell wall content), this polymer remains essential for the cell wall, guaranteeing its integrity. Inhibition of chitin synthases by nikkomycin Z in S. monoica causes cell lysis through tip bursting. This indicates that chitin most likely substitutes cellulose at the tip and plays a key role in tip growth (Guerriero et al., 2010). The chitin detected in S. monoica corresponds to crystalline α-chitin and, depending on the physiological state of the oomycete cell, it occurs as granular structures in hyphal cell walls or it is deposited as microfibrils in regenerating protoplasts (Gay et al. 1992; Bulone et al. 1992). Mélida et al. (2013) proposed a model distinguishing three types of oomycete cell walls (Figure 4), based primarily on the content of N-acetylglucosamine (GlcNAc), the monomer of chitin. Members of the Peronosporales order, including P. infestans, have type I cell walls, lacking GlcNAc. Another unique feature for this type of cell wall is the presence of glucuronic acid and higher contents of mannose. In contrast, the Saprolegniales order, which comprises species like Saprolegnia, Achlya and Leptolegnia sp., is characterized by type II cell walls with a GlcNAc content up to 5%. Moreover, type II cell wall are distinguished from type I and type III by the occurrence of 1,3,4-linked glucosyl residues in the cell wall alkali-insoluble fraction. This suggests the presence of cross-links between cellulose and 1,3-glucans, which have never been described before. The type III cell wall contains the highest amount of GlcNAc (>5%). Type III is represented by the plant pathogen Aphanomyces euteiches. Interestingly, this type of cell wall, in addition to containing 1,4-linked GlcNAc residues characteristic of chitin, contains 1,6-linked GlcNAc residues. It is the first time that this type of linkage has been shown to occur in eukaryotic microorganisms. The detailed cell wall analysis by Mélida et al. (2013) revealed that the 1,4-linked GlcNAc residues are most likely derived from chitin of a low degree of polymerization, which is contradictory to previous studies on A. euteiches cell walls where no chitin was detected (Badreddine et al., 2008). Additionally, type III and I cell walls have higher contents of 1,3,6- glucose, which indicates the presence of branched 1,3-glucans at the C-6 positions of the main chain (Mélida et al., 2013).

Fig. 4. Composition of the three different cell wall types of representatives species of the Saprolegniales and Peronosporales orders (bold font) (from Mélida et al. 2013).

Fungal cell walls

The most studied cell walls in eukaryotic microorganisms are from yeast and fungi. Cell wall composition varies significantly between different species within the kingdom Fungi (Table 2), but all exhibit structural similarities. Cell wall components occur as two layers, the inner layer being essentially alkali insoluble and the outer layer being alkali soluble (Latgé and Calderone, 2006; Latgé, 2007). The inner layer is considered as a structural skeleton of the cell wall and in yeast it consists largely of β- 1,3-glucans and chitin. The outer layer is composed of mannans and proteins, that are linked to the inner wall via β-1,6-glucans (Gow et al., 2017) (Figure 5).

Fig. 5. Cell wall model of the yeast Candida albicans (from Gow et al., 2012; Gow et al., 2017).

β-1,3-Glucan is the major component of yeast and fungal cell walls and constitutes from 30 to 80% of the cell wall mass (Free, 2013). Chains of β- 1,3-glucan are generally branched with β-1,6 linkages and form a glucan scaffold to which various cell wall components like chitin, β-1,6-glucans and some mannoproteins are covalently attached (Bowman and Free, 2006; Klis et al., 2006).

Chitin is another structurally important constituent that contributes to the rigidity, strength and integrity of the cell wall. It accounts for 1 to 2% of the yeast cell wall mass, while filamentous fungi like Neurospora and Aspergillus contain between 10 to 20% of the polymer (Table 2) (Bowman and Free, 2006; Free, 2013).

In yeast species, β-1,6-glucan represents 10 to 15% of the cell wall, but this polymer is absent in many filamentous fungi including Neurospora and Aspergillus (Lipke and Ovalle, 1998; Klis et al., 2006; Free, 2013). β- 1,6-Glucan is a highly branched polysaccharide, which functions as a linker connecting β-1,3-glucans and chitin with cell wall glycoproteins (Kollár et al., 1997). In contrast, cell walls of Aspergillus and Neurospora contain α-1-3-glucans, which are not detected in S. cerevisiae and C. albicans. Moreover, A. fumigatus contains galactomannans and mixed- linkage glucans that consists of both β-1-3 and β-1-4 linkages covalently

10 bound to the cell wall core (Table 2) (Bernard and Latgé, 2001; Bowman and Free, 2006).

Table 2. Cell wall composition in two examples of yeast and fungal species: Saccharomyces cerevisiae and Aspergillus fumigatus (modified from Free, 2013 and Klis et al., 2006)

Cell wall component S. cerevisiae A. fumigatus Chitin Yes (1-2%) Yes (10-20%) β-1,3-Glucan Yes (50-55%) Yes (20-35%) β-1,6-Glucan Yes (10-15%) No Mixed β-1,3/1,4-glucan No Yes α-1,3-Glucan No Yes (35-46%) Mannoproteins Yes (30-50%) No Galactomannan No Yes (20-25%)

Cell wall proteins are divided into wall-associated enzymes (WAE) and “structural” proteins. The former have enzymatic activity and are involved in cell wall synthesis and remodeling. This group includes hydrolases (e.g. chitinases, glucanases) or transglycosidases (e.g. β-1,3- glucanosyltransferase) (Mouyna et al., 2000). “Structural” proteins are nonenzymatic proteins participating in cell adhesion, fusion or mating (Bowman and Free, 2006). Glycoproteins constitute the majority of the proteins associated with the cell wall. In yeast these are N- and O-glycosylated mannoproteins, which make up between 30-50% of the S. cerevisiae and C. albicans cell walls (Bowman and Free, 2006; Free, 2013). A number of glycoproteins are covalently bound to the glycan network. Based on their mode of binding, the proteins can be distinguished between glycosylphosphatidylinositol (GPI)-proteins, mild alkali- releasable proteins and disulfide-linked proteins. Most of the yeast mannoproteins are GPI cell wall proteins (GPI-CWP) that are crosslinked to glucans via the GPI remnant (a part of GPI anchor left after cleavage of the lipid moiety). These GPI-CWP are crosslinked to β-1,6- glucans, which in turn are attached to β-1,3-glucans and chitin. A minor group of alkali-soluble proteins are directly ester-linked to β-1,3-glucans. In addition, there are various proteins that are disulfide bonded to other

CWP (Lipke and Ovalle, 1998; Yin et al., 2005; Klis et al., 2006; Orlean, 2012).

Chitin

Chitin is one of the most abundant biopolymers in nature. It is present in many organisms including fungi, some oomycetes and arthropods such as crustaceans and insects (Hudson and Smith, 1998; Mélida et al. 2013). Chitin is a linear polymer of β-1,4-linked N-acetylglucosamine (GlcNAc) residues (Figure 6). It forms crystalline microfibrils stabilized by intermolecular hydrogen bonds between the N-H groups of one chain and the C=O groups of a neighboring chain as well as a number of hydrogen bonds between hydroxyl (O-H) and carbonyl groups (C=O) (Cabib, 1981; Minke and Blackwell; 1978).

Fig. 6. Chitin chain composed of repeated β-1,4-N-acetylglucosamine units. Every monomer is flipped by 180° with respect to its adjacent monosaccharide.

Inter- and intramolecular hydrogen bonding, as well as the intersheet bonding, makes chitin hydrophobic and insoluble in water and most organic solvents (Cabib, 1981; Zargar et al. 2014). Based on the organization of chains, chitin occurs as three different forms called allomorphs α, β and γ (Figure 7) (Cabib, 1981; Hudson and Smith, 1998).

Fig. 7. Representation of the three crystalline forms (allomorphs) of chitin: α, β and γ (redrawn from Anitha et al., 2014).

α-Chitin is the most common form in nature and can be found in the cell wall of fungi and the exoskeleton of crustaceans like crabs or shrimps. In α-chitin, the GlcNAc chains are oriented antiparallel to one another, allowing very strong hydrogen interactions and creating the most compact structure of chitin (Minke and Blackwell, 1978). β-Chitin is composed of chains oriented into the same direction (parallel). This form of chitin is far less abundant than α-chitin, and mainly occurs in squid pens (Loligo) and diatom spines (Lavall et al., 2007; Blackwell, 1969). β- Chitin is also less stable, since it lacks intersheet hydrogen bonds (Hudson and Smith, 1998). The third allomorph, γ-chitin, is a rare form of chitin that has been reported in several insects and Loligo stomach (Hudson and Smith, 1998). It has not been completely characterized, but it was proposed that the repeating structure of γ-chitin consists of two adjacent chains that are oriented parallel, and a third chain that runs in the opposite direction (Rudall, 1963).

Cellulose

Cellulose is a linear polymer that consists of glucose units connected by β- 1-4 linkages (Figure 8). It is the most abundant polymer on earth, produced by a large diversity of organisms including plants, oomycetes, algae, tunicates and some bacteria. In plants, depending on the source and extraction methods, cellulose chains typically contain between 500 and 15000 glucose units (Somerville, 2006).

Fig. 8. Molecular structure of cellulose. Every glucose residue is rotated 180o with respect to its neighbor, making cellobiose a suggested repeating unit of cellulose.

Cellulose, similarly to chitin, is hardly soluble in any solvent due to the formation of many inter- and intramolecular hydrogen bonding (O-H-O), and numerous (C-O--H) hydrogen bonds between neighboring sheets. This structure confers rigidity to cellulose and is responsible for its insolubility (Jarvis, 2003). In addition to crystalline segments, cellulose microfibrils contain amorphous, less organized areas (Figure 9).

Fig. 9. Cellulose micofibrils containing both crystalline and disorganized regions (adapted from Moon et al., 2011).

X-ray diffraction revealed that crystalline cellulose exists as four different allomorphs: cellulose I, II, III and IV, which can be distinguished by their specific unit cell geometry (Moon et al., 2011). Cellulose I is the native form of cellulose divided into cellulose Iα and Iβ. Both forms co-exist with each other in nature, where cellulose Iα is predominantly found in algae, e.g. Valonia ventricosa, and bacteria such as Gluconacetobacter xylinus (recently renamed Komagataeibacter xylinus), while cellulose Iβ is the main form of cellulose in higher plants (e.g. wood, cotton) (Atalla and Vanderhart, 1984). Cellulose I consists of parallel chains, with reducing ends pointing in the same direction. In contrast, cellulose II

14 consists of chains organized in an anti-parallel fashion. This type of cellulose is naturally produced by the alga Halicystis (Sisson, 1938) and some mutant strains of Gluconacetobacter xylinus (Kuga et al., 1993) but it can also be obtained by chemical treatment, e.g. by alkali (NaOH) treatment of cellulose I. The alkali treatment causes swelling and disorganization of chains, which after washing rearrange in the thermodynamically more favorable anti-parallel fashion (Klemm et al., 2005). Due to its silky texture, cellulose II is widely used in the textile industry. Type III cellulose can be obtained by chemical treatment of cellulose I or cellulose II. Cellulose IV is a natural form of cellulose found in oomycetes and plant cell walls (Chanzy et al., 1979; Helbert et al., 1997; Bulone et al., 1992), and is considered a disordered form of cellulose I (Wada et al., 2004).

β-1,3-Glucans

As mentioned above, β-1,3-glucans are the main constituents of fungal cell walls. The approximate degree of polymerization of glucan chains is around 1500 (Klis et al., 2002). They are composed of repeating glucose units connected by β-1,3-linkages. As opposed to plant β-1,3-glucan (callose), which is linear, β-1,3-glucans in oomycete and fungal species are branched at the C6 position (Figure 10). These branched polymers are assembled into a three-dimensional network, stabilized by hydrogen bonding between glucan chains (Klis et al., 2006; Latgé, 2007). They have a low level of crystallinity due to the presence of the side-chains, which are typically irregularly distributed along the main chain and prevent crystallization (Klis et al., 2002).

Fig. 10. Scheme representing the general structure of a β-1,3-glucan with a branching point at the C6 position.

Polysaccharide biosynthesis

Glycosyltransferases

Glycosyltransferases (GT) catalyze the transfer of glycosyl residues from an activated donor (usually a nucleoside-diphospho-sugar, e.g. UDP- glucose, UDP-N-acetylglucosamine) to a specific acceptor molecule, creating glycosidic bonds (Saxena et al., 1995). A large number of different GTs are involved in the biosynthesis of disaccharides, oligosaccharides, polysaccharides as well as in the synthesis of glycoproteins and glycolipids. At present, they are classified in 106 different families in the CAZy (carbohydrate-active enzyme) database (www.cazy.org) based on sequence similarity, 3-dimensional structure, when available, and reaction mechanism (Campbell et al., 1997; Coutinho et al., 2003). GTs occur as two types of enzymes: they act either as processive enzymes, able to catalyze the transfer of multiple sugar residues to a growing carbohydrate chain, or non-processive enzymes, transferring single sugar moieties to the acceptor (Saxena et al., 1995).

Both cellulose (CesA) and chitin (CHS) synthases are processive enzymes and belong to GT family 2.

Catalytic mechanisms

Based on their catalytic mechanism, GTs are classified as inverting or retaining enzymes (Figure 11). In the inverting mechanism, the configuration of the anomeric carbon of the product is changed with respect to the sugar donor (from α to β configuration), whereas in the retaining mechanism it remains unchanged (α configuration) (Lairson et al., 2008).

Fig. 11. Scheme representing the inverting and retaining mechanisms of GTs (from Lairson et al., 2008).

All GTs from family 2, including CHS and CesA enzymes, use an inverting mechanism of catalysis. This involves a one-step nucleophilic substitution reaction, in which at least two catalytic carboxylates are involved (Figure 12). The first one functions as a general base during catalysis. It activates the acceptor sugar by deprotonating the hydroxyl group at the non- reducing end of the molecule. The activated acceptor is then able to perform a nucleophilic attack on the anomeric carbon of the α-linked NDP-sugar (e.g., UDP-GlcNAc, UDP-Glc) which results in the formation

17 of β-linked products (e.g., chitin, cellulose). The other carboxylic residue(s) assists in removing the nucleotide group, which additionally is stabilized by metal ions like Mn2+ or Mg2+ (Figure 12; Charnock et al., 2001; Saxena et al., 1995; McNamara et al., 2015).

Fig. 12. Inverting reaction mechanism of GTs (from Lairson et al., 2008 and Charnock et al., 2001).

The retaining mechanism of GTs involves a two-step reaction (Figure 13). This reaction is initiated by a nucleophilic attack of a carboxyl residue on the anomeric carbon of the α-linked sugar donor leading to the formation of an enzyme-donor intermediate in β configuration. In the second step, the leaving phosphate group most likely acts as a catalytic base which activates the hydroxyl group of an acceptor sugar that performs a nucleophilic attack on the enzyme-donor intermediate, consequently producing an α-linked product (Lairson et al., 2008) (Figure 13).

Fig. 13. Retaining reaction mechanism of GTs (from Lairson et al., 2008).

Chitin synthases

Chitin is synthesized by membrane-bound proteins called chitin synthases (CHS). These are processive enzymes which utilize UDP- GlcNAc as a substrate and elongate chitin chains from their non-reducing ends (Kamst et al., 1999). However, the detailed mechanism of chitin biosynthesis, initiation of polymerization and assembly process into microfibrils remains unknown.

Yeast and fungal chitin synthases

Different numbers of Chs genes have been identified and studied in various yeast and fungal species. In S. cerevisiae and C. albicans three and four genes, respectively, were identified to encode chitin synthases (Bulawa et al., 1986; Sburlati and Cabib, 1986; Valdivieso et al., 1991; Munro and Gow, 2001; Munro et al., 2003). Three CHS isoforms from S. cerevisiae, ScCHS1, ScCHS2 and ScCHS3 have been very well characterized and were found to have different biological functions and biochemical properties. ScCHS1 is involved in cell wall reparation (Cabib et al., 1989), ScCHS2 in the formation of the septum prior to cell division (Shaw et al., 1991; Silverman et al., 1988) and ScCHS3 in the synthesis of more than 90% of the cell wall chitin, which includes the chitin ring

19 formed between the mother and daughter cells and chitin in the lateral cell wall (Shaw et al., 1991). In S. cerevisiae the mutation of each individual gene does not affect cell viability, but results in slower growth and morphological abnormalities (Shaw et al., 1991). The double mutant Δchs2Δchs3, however, is lethal for the cell (Shaw et al., 1991). In contrast, the mutation of Chs1 in C. albicans is lethal, demonstrating that this gene is vital for the microorganism (Munro et al., 2001). This shows different functions, requirements and significance of chitin synthases between species.

In filamentous fungi, on the other hand, the number of Chs genes is usually higher, which is also reflected in a higher chitin content in the cell wall (15-20%) than in budding yeasts (~2%). Seven genes were identified in Magnaporthe oryzae (Kong et al., 2012) and N. crassa (Borkovich et al., 2004), eight in A. fumigatus (Mellado et al., 2003; Walker et al., 2015) and up to 11 in Aspergillus oryzae (Latgé and Calderon, 2006). The occurrence of multiple chitin synthase genes is correlated with chitin synthesis at different developmental stages of the fungal cells, where they play specific roles, e.g. in appressoria and septa formation, pathogenesis, hyphal growth or conidiogenesis. Some of the genes are functionally redundant (Kong et al., 2012; Liu et al., 2016). In most filamentous fungi, mutational analysis of single Chs genes revealed that individual genes are generally not essential for cell survival (Mellado et al., 2003; Kong et al., 2012). The disruption of the individual Chs genes is most likely compensated by the activity of the remaining chitin synthases or increased synthesis of other cell wall carbohydrates, like glucans (Mellado et al., 2003). Typically, the mutation of major Chs in filamentous fungi results in slower hyphal growth, swollen tips and increased hyphal branching (Mellado et al., 1996; 2003; Borgia et al., 1996). Similar morphologies were observed in S. monoica and S. parasitica mycelium in the presence of the chitin synthase inhibitor nikkomycin Z (Guerriero et al., 2010; Rzeszutek et al., 2019a). The results in these species indicated that the activity of CHS essentially occurs in the apex of fungal and oomycete cells. They are consistent with the results obtained by McMourrough et al. (1971), who, using the radioactive substrate UDP- [3H]GlcNAc, demonstrated that chitin biosynthesis occurs mainly in the apical region of the mycelial wall of Mucor rouxii. Moreover, apical localization of fluorescently labeled CHSs was observed in the growing

20 hyphae of N. crassa (Riquelme et al., 2007) and Ustilago maydis (Schuster et al., 2016).

Oomycete chitin synthases

Oomycete CHSs have not been studied as extensively as their fungal counterpart, but putative Chs genes have been found in all analyzed species regardless of the presence of chitin in the cell wall (Klinter et al., 2019). The genomes of Peronosporales such as P. infestans, Phytophthora sojae or Phytophthora ramorum usually carry one or two Chs genes (Haas et al., 2009; Tyler at al., 2006), while Saprolegniales like S. parasitica, carry up to six Chs genes with a chitin content of around 2% (Jiang et al., 2013; Rzeszutek et al. 2019a, Klinter et al., 2019). In oomycetes, CHSs together with other cell wall polysaccharide synthases were shown to be located in detergent-resistant domains of the plasma membrane (Briolay et al., 2009). To date, the catalytic activity of only two CHSs, namely SmCHS2 and SpCHS5, was demonstrated to have a N-acetylglucosaminyl transferase activity, which led to the production of crystalline chitin in vitro (Guerriero et al., 2010; Rzeszutek et al., 2019b). Two chitin synthases were also identified in A. euteiches (Baddredine et al., 2008), whose cell wall analysis interestingly revealed the greatest amount of GlcNAc-based carbohydrates in all characterized oomycete cell walls (~10%). The data indicated the participation of A. euteiches Chs genes in the synthesis of either chitin with a low degree of polymerization or soluble oligosaccharides that can be linked to other cell wall glycans (Mélida et al., 2013; Badreddine et al., 2008). Some oomycete species, however, like P. infestans, do not contain chitin in their cell wall despite the presence of putative Chs genes. The function of the single P. infestans Chs gene has never been determined. Nonetheless, recent studies suggest that the activity of the corresponding protein is required for the vegetative growth of this species. Moreover, the particularly high expression of this putative Chs during plant infection indicates a potential specific role of this gene in pathogenesis (Hinkel et al., 2017).

Domains of chitin synthases

All known CHSs carry a conserved set of motifs: D, D (domain A), involved in substrate binding, and D, QXXRW (domain B), involved in acceptor binding (Saxena et al., 1995; 2001; Charnock et al., 2001). Both domains A and B are found in most processive GTs, while non-processive GTs contain domain A only (Saxena et al., 1995). In fungal and oomycete CHSs, these motifs have the following sequences: D, Dx(D/G), EDR and Q(R/Q)XRW (Choquer et al., 2001; Rzeszutek et al., 2019a) (Figure 14). Mutational analysis of these invariant residues in ScCHS2 revealed their significance and critical role for CHS activity (Nagahashi et al., 1995). Another characteristic of CHSs is the presence of a Con2 domain (second conserved region) at the C-terminal end of the proteins that was characterized in ScCHS2. Since the mutations of amino acids in this region resulted in the production of oligomers not longer than chitobiose, it was suggested that Con2 is involved in enzyme processivity (Yabe et al., 1998). Even though CHSs share a common domain organization, slight differences exist in their active sites, which lead to different susceptibility to inhibitors. For example, ScCHS2 and ScCHS1 are inhibited by the chitin synthase inhibitor nikkomycin Z (NZ), while ScCHS3 is barely affected (Cabib, 1991; Gaughran et al., 1994). Interestingly, all three corresponding enzymes from C. albicans, namely CHS1, CHS2 and CHS3 were sensitive to NZ (Kim et al., 2002).

Fig. 14. Topology model of S. parasitica CHS5 with its conserved motifs.

All CHSs have multiple transmembrane domains (TM) located mainly at their C-terminal ends. For example there are eight C-terminal TM domains predicted for S. parasitica CHS5 (Figure 14). Interestingly, several CHSs from S. parasitica and S. monoica contain MIT (Microtubule Interacting and Trafficking) domains at their N-terminal ends (Guerriero et al., 2010; Rzeszutek et al., 2019a). This domain plays a role in intracellular protein trafficking in many species (Cicciarelli et al., 2003), however oomycete CHSs are the first known carbohydrate-active enzymes where this domain was identified (Guerriero et al., 2010). In many fungi, including M. rouxii, S. cerevisiae and N. crassa, small vesicular structures called chitosomes were identified to be involved in CHS delivery to the growth (tip) region of the cells (Bracker et al., 1976; Bartnicki Garcia et al., 1978). These structures, however, have never been found in oomycetes (Leal-Morales et al., 1997), and therefore it was suggested that the MIT domain of oomycete CHS is involved in the intracellular transport and/or regulation of the enzymes instead of chitosomes (Brown et al., 2016).

Regulation of chitin synthases

The formation of chitin occurs mainly in areas of growth and cell wall remodeling (Cabib et al., 2001, Lenardon et al., 2007). Thus, CHS activity has to be strictly regulated at specific times and locations. Many different yeast and fungal CHSs are of zymogenic nature and were shown to be proteolytically regulated in vitro (Sburlati and Cabib, 1986; Gay et al., 1989; Martínez-Rucobo et al., 2009). In the case of the CHSs from S. cerevisiae, only CHS1 (Duran and Cabib, 1978; Orlean, 1987) and CHS2 are highly activated by trypsin (Sburlati and Cabib, 1986; Uchida et al., 1996; Martínez-Rucobo et al., 2009). In contrast, the activity of ScCHS3 is partially reduced when treated with this protease (Orlean, 1987). Interestingly, ScCHS2 shows significant activity in vitro also without any protease treatment, indicating that both forms, i.e. the full size and processed protein, can act as fully active enzymes (Uchida et al., 1996). A stimulatory effect of trypsin was also demonstrated in vitro for the oomycete chitin synthases SmCHS2 and SpCHS5 (Guerriero et al., 2010; Rzeszutek et al., 2019b). Nonetheless, it remains elusive whether the proteolytic enzymes act directly or indirectly on chitin synthases. Trypsin

23 was found to activate ScCHS2 indirectly by acting on other soluble proteases, which subsequently activate CHS (Martínez-Rucobo et al., 2009). On the other hand, purified SpCHS5 seems to be activated directly in the presence of trypsin (Rzeszutek et al., 2019b). Proteolytic activation of CHS in vitro may reflect chitin synthase regulation in vivo. However, no processed forms of the enzymes have been identified in S. cerevisiae and Saprolegnia species. Chitin synthase activity was also shown to be stimulated in vitro by free GlcNAc (Sburlati and Cabib, 1986; Orlean, 1987; Keller and Cabib, 1971; McMurrough and Bartnicki-Garcia, 1971). The exact role of GlcNAc is not known, but it was proposed to serve as a primer or allosteric activator of the enzyme. Free GlcNAc was demonstrated to serve as a primer for the synthesis of chitin oligosaccharides by NodC (a bacterial chitin synthase homologue with low processivity) (Kamst et al., 1999) and GlcNAc analogs were also shown to prime the formation of chitooligosaccharides by ScCHS2, suggesting that free GlcNAc acts in a similar way (Gyore et al., 2014). However, in several CHSs, GlcNAc was reported to function as an allosteric activator (McMurrough and Bartnicki-Garcia, 1971; Horsch et al., 1996), possibly representing another regulatory mechanism of chitin biosynthesis. In addition to GlcNAc, longer chitooligosaccharides were proposed to act as primers for the addition of GlcNAc residues (Becker at al., 2011). These data, however, were from in vitro studies, which do not necessarily mean that GlcNAc or chitooligosaccharides act as primers in vivo. Another regulatory mechanism of proteins is phosphorylation. This process was shown to be required for the correct localization and function of C. albicans CHS3 (Lenardon et al., 2010). Phosphorylation sites were detected at the N-terminal end of ScCHS2, but their role is not clear. Nonetheless, dephosphorylation of ScCHS2 was accompanied by a loss of enzyme stability, in particular a higher sensitivity to proteases (Martinez- Rucobo et al., 2009). These observations revealed a potentially important role of phosphorylation and dephosphorylation processes in CHS localization and stability.

Cellulose synthases

Cellulose synthases (CesA) are GTs that utilize UDP-activated glucose to form β-1-4-glucans. UDP-Glucose can be synthesized either by a pyrophosphorylase from UTP and glucose-1-phosphate or by sucrose synthase (SuSy) from UDP and sucrose (Figure 15). A membrane-bound form of SuSy was proposed to be associated with the cellulose synthase machinery (Amor et al., 1995; Fujii et al., 2010), providing substrate directly to the enzyme. However, the existence of direct interactions between SuSy and CesA proteins has never been confirmed.

Fig. 15. Model representing cellulose biosynthesis in plants (adapted from McFarlane et al., 2014). SuSy: sucrose synthase; CSC: cellulose synthase complex.

In the bacterium Gluconacetobacter xylinus (Koyama et al., 1997) and the blackberry Rubus fruticosus (Lai Kee Him et al., 2002) it was demonstrated that cellulose chains are elongated from their non-reducing ends. These data were confirmed from the crystal structure of the bacterial cellulose synthase (Bcs) from Rhodobacter (Morgan et al., 2013), which additionally revealed the involvement of a single binding site for the substrate and transfer of one glucose unit at a time. The mechanism of initiation of the elongation process is however still not known. In vitro studies suggested sitosterol-β-glucoside functions as a

25 primer for cellulose biosynthesis (Peng et al., 2002). However, this has never been demonstrated in vivo. Moreover, recent studies on the bacterial cellulose synthase complex (BcsA-B) showed that cellulose elongation occurs directly from UDP-Glc in the absence of any lipid- linked oligosaccharide (Omadjela et al., 2013), which challenges the concept that sitosterol-β-glucoside acts as a primer.

Cellulose synthases are arranged in large complexes that are located in the plasma membrane. These enzyme complexes were imaged for the first time in the green alga Oocystis. Because of their occurrence at the end of growing cellulose microfibrils, they were called terminal complexes (TC) (Brown and Montezinos, 1976). Depending on the organism, TC can be organized in different ways. In plants, but also some green algae, the cellulose synthase subunits of the TC are organized in rosettes, while in bacteria, tunicates and most algae they are assembled in linear arrays (Figure 16a) (Mueller and Brown, 1980; Brown, 1996). None of these structures, however, have been identified in oomycetes.

Fig. 16. Scheme showing the organization of rosette and linear types of terminal complexes (adapted from Brown, 1996) (a). Model representing the structure of a rosette containing three CesA molecules in each rosette subunit, and producing elementary microfibrils composed of 18 cellulose chains each (adapted from McFarlane et al., 2014) (b).

The shape and size of the cellulose microfibrils depend on the configuration of the subunits in the complexes and the number of catalytic proteins within the TC. Plant rosettes consist of six subunits, which each contain multiple catalytic CesA proteins. The presence of catalytic CesA subunits in rosettes was demonstrated by freeze-fracture electron microscopy combined with immunogold labeling (Kimura et al., 1999). Earlier models proposed that each of the six rosette subunits contains six CesA proteins producing a total of 36 cellulose chains per rosette and microfibril (Somerville, 2006; Doblin, et al., 2002). The individual chains produced by the rosettes self-assemble into microfibrils, which further crystallize into larger fibrillar structures. The model of 36 chains per rosette is mainly based on the TC structure and is consistent with the dimensions of individual microfibrils, which have a lateral size of 2-5 nm (Ha et al., 1998; Guerriero et al., 2010). However, more recent studies based on improved imaging and in silico modeling as well as scattering and spectroscopic techniques, revealed that plant rosettes most likely contain three CesA proteins in each of the six subunits, consequently producing elementary fibrils composed of 18 instead of 36 chains (Figure 16b) (Newman et al., 2013; Nixon et al., 2016).

In Arabidopsis, which contains 10 CesA genes, CesAs are organized in two different complexes each composed of at least three different CesA isoforms. One of the complexes is involved in primary cell wall formation and consists of CesA1, 3 and 6, where CesA6 can be replaced by CesA2, 5 or 9 without loss of function. The second complex is involved in secondary cell wall formation and contains CesA4, 7 and 8 (Arioli et al., 1998; Taylor et al., 2000, 2003; Desprez et al., 2007; Persson et al., 2007). The organization of these proteins within the complex is however not known. Recent studies showed that the single catalytic subunits CesA8 and CesA5 from the hybrid aspen Populus tremula x tremuloides and the moss Physcomitrella patens, respectively, are sufficient to produce cellulose microfibrils without being in a complex with any other CesA isoforms (Purushotham et al., 2016; Cho et al., 2017). This indicates that CesA5 and CesA8 are able to form homooligomers, most likely homotrimers, as also reported for Arabidopsis CesA1 (Vandavasi et al., 2016). The formation of trimers is further consistent with the model that

27 proposes the presence of three CesA molecules per subunit of the rosette (Nixon et al. 2016). In Arabidopsis, additional proteins like KORRIGAN (KOR), COBRA (COB) and KOBITO1 (KOB1) were shown to be involved in cellulose formation. KORRIGAN, a membrane-bound endo-β-1,4-glucanase, is part of the cellulose synthase complex and is required for cellulose synthesis (Figure 15) (Vain et al., 2014). Mutation of the KOR1 gene resulted in reduced growth and decreased content of cellulose in the cell wall (Nicole et al., 1998; Sato et al., 2001), but the exact function of the protein in cellulose biosynthesis remains unknown. COBRA, a GPI- anchored protein, and the membrane-bound protein KOBITO1, are also needed for normal cellulose formation. Mutations of the KOB1 and COB genes affect plant growth and cause cellulose deficiency (Pagant et al., 2002; Roudier et al., 2005).

The bacterial cellulose synthase (Bcs) complex is encoded by an operon that carries multiple genes, i.e. bscA, B, C and Z (Römling et al., 2002; McNamara et al., 2015). In Rhodobacter sphaeroides it was demonstrated that a complex composed of two different protein subunits, BcsA and BcsB, is sufficient for in vitro cellulose synthesis (Omadjela et al., 2013). BcsA is responsible for the catalytic activity, but its activity strictly depends on the presence of BcsB, which is a periplasmic protein anchored in the plasma membrane. The non-catalytic BcsB protein contains a carbohydrate-binding domain (CBD), which most likely helps in glucan translocation across the periplasm. Moreover, the presence of the signaling molecule cyclic diguanylate (cyclic-di-GMP) is essential for Bcs activity (Omadjela et al., 2013; Morgan et al., 2014). This small molecule works as an allosteric activator, binding to the PilZ domain located at the C-terminal end of the BcsA subunit (Amikam et al., 2006). The interaction of c-di-GMP with the PilZ domain induces a movement of the gating loop that blocks the active site, consequently allowing UDP-Glc to bind to the active site of the catalytic subunit.

Oomycete cellulose synthases

Even though cellulose is the main component of oomycete cell walls, the mechanism of cellulose biosynthesis in these species is largely unknown. The presence of four cellulose synthase genes was discovered in Peronosporales species like P. infestans, P. sojae, P. ramorum and Plasmopara viticola (Grenville-Briggs et al., 2008; Blum et al., 2010), as well as in Pythiales species such as Pythium aphanidermatum (Blum and Gisi, 2012). In P. infestans, the function of the CesA genes in cellulose synthesis was investigated by silencing all four genes. This resulted in reduced cellulose content in the cell wall and in the lack of ability to form proper appressoria, which are infection structures required for penetration of the plant tissues by the pathogen. Moreover, the inhibitor of cellulose biosynthesis, 2,6-dichlorobenzonitrile (DCB), causes total loss of pathogenicity, indicating that cellulose synthase genes are important for the infection process (Grenville-Briggs et al., 2008). In the Saprolegniales S. monoica, three CesA genes were identified. Slower mycelial growth, higher expression of CesA genes and elevated in vitro CesA activity in the presence of cellulose synthase inhibitors in the culture medium also point to a role of these genes in cellulose formation (Fugelstad et al., 2009). The presence of six CesA genes and production of cellulose in vitro by enzyme extracts were demonstrated for S. parasitica (Rzeszutek et al., 2019c). However, to date, none of the individual oomycete cellulose synthases have been functionally characterized.

Domains of cellulose synthases

As mentioned earlier, processive GTs, including cellulose synthases, contain three conserved aspartic residues (D, D, D) and the pentapeptide QXXRW (Campbell et al., 1997; Saxena et al., 1995; 1997). In all described CesAs, the motifs containing key aspartic residues are DDG, DxD and TED. The crystallographic structure of the bacterial cellulose synthase from R. sphaeroides (RsBsc) revealed that the first Asp residues in the DDG and DxD motifs mediate substrate binding, while the TED and QXXRW motifs are involved in the binding of the glucan (glucose) acceptor (Morgan et al., 2013; McNamara et al., 2015). The TED motif is part of a zinc finger helix involved in glucan translocation, and its aspartic

29 acid residue probably functions as the general base during catalysis (Morgan et al., 2013; 2016; McNamara et al., 2015). These conserved catalytic domains in CesA proteins are flanked by multiple transmembrane regions, which form a channel for translocation of the growing cellulose chains across the membrane to the extracellular space (Morgan et al., 2013). Interestingly, two groups of oomycete cellulose synthases, classified as CesA2 (recently CesA1 from Phytophthora spp. reclassified as CesA2; Rzeszutek et al. 2019c) and CesA4, contain plekstrin homology (PH) domains at their N-terminal ends (Fugelstad et al., 2009; Grenville-Briggs et al., 2008). PH domains have been shown to be present in eukaryotic proteins involved in signaling pathways and they are known to be involved in targeting proteins to different membrane compartments (Lemmon et al., 1996). To date, oomycete CesAs are the first carbohydrate-active enzymes known that have been shown to contain this type of domain. The functional characterization of the PH domain from SmCesA2 revealed its role in the trafficking and/or targeting of the protein (Fugelstad et al., 2012). This is possibly a similar role as that attributed to the MIT domain of CHSs. This suggests that the PH domain is functionally different from the zinc finger domain found at the N-terminus of plant CesAs, which is involved in protein dimerization (Kurek et al., 2002) possibly supporting the formation of the rosette complexes in plants. In contrast to plant and oomycete cellulose synthases, the bacterial BcsA contains a PilZ domain involved in the binding of the c-di-GMP activator.

β-1,3-Glucan synthase

β-1,3-Glucan synthase (GS) is responsible for the formation of β-1,3- glucans. It is a processive membrane-bound protein that belongs to the GT family 48. The enzyme uses UDP-glucose as a substrate, which is synthesized at the cytosolic side of the plasma membrane. During the synthesis, the enzyme extrudes the growing glucan chain across the membrane into the extracellular cell wall space (Douglas, 2001; Latgé and Calderone, 2006).

In S. cerevisiae, two genes encoding two catalytic subunits of the β-1,3- glucan synthase have been identified, namely FKS1 and FKS2. Individual

30 mutations of either the FKS1 or FKS2 gene are not lethal, however FKS1 mutants show slower growth and cell wall alterations. On the other hand, the simultaneous disruption of both FKS1 and FKS2 leads to cell death (Douglas et al., 1994; Mazur et al., 1995). Filamentous fungi, including A. fumigatus, contain a single β-1,3-glucan synthase, FKS1, which was also shown to be crucial for cell viability (Beauvais et al., 2001). This shows how essential these genes and synthesis of β-1,3-glucans are for cell survival. Currently, one of the β-1,3-glucan synthase inhibitors, caspofungin, is clinically used as a therapeutic agent against candidiasis and aspergillosis. In the fungus U. maydis, GS was shown to be co-delivered to the plasma membrane in the same secretory vesicle as CHSs (Schuster et al., 2016). The activity of both enzymes is coordinated during cell wall synthesis.

In the oomycete S. monoica, GS activity has been localized in detergent- resistant microdomains of the plasma membrane (Briolay et al., 2009). The activity of the GS from S. monoica and S. parasitica was demonstrated in vitro using mycelial extracts (Bulone et al., 1990; Fèvre and Rougier, 1981; Rzeszutek et al., 2019c), but the corresponding genes have never been directly linked to enzyme activity. To date, only one oomycete protein involved in β-1,3-glucan synthesis has been identified, and it is a regulatory protein from the annexin family (Bouzenzana et al., 2006).

Cell wall biosynthetic enzymes as potential targets for anti-oomycete agents

Cell wall related inhibitors

Cell wall formation is a crucial process for microorganisms, and, in eukaryotic species, it involves many different enzymes including cellulose, chitin and β-1,3 glucan synthases. The studies discussed in the previous sections highlight the importance of these enzymes in the growth and cell wall integrity of fungal and oomycete species. Thus, gaining insight into cell wall biosynthesis mechanisms is of high

31 importance to target specific cell wall synthesizing enzymes for disease control. There are several inhibitors that were shown to effectively affect cell wall formation. These include nikkomycin Z, DCB, echinocandins, compound I, CGA and flupoxam, described below. Moreover, some of these specific chemical inhibitors are useful tools to study the enzymes involved in cell wall biosynthesis.

Nikkomycin Z

Nikkomycin Z (NZ) is a nucleoside-peptide antibiotic produced by the bacterium Streptomyces tandae and has been shown to be a specific inhibitor of yeast, fungal and oomycete CHSs (Cabib, 1991; Gaughran et al., 1994; Guerriero et al., 2010; Kim et al., 2002; Gow and Selitrennikoff, 1984; Rzeszutek et al., 2019b). The structure of NZ mimics that of the CHS substrate (UDP-GlcNAc) and consequently acts as a competitive inhibitor of CHS (Figure 17) (Gooday, 1990). In S. monoica and S. parasitica, NZ strongly inhibits CHS activity in vitro and causes a reduced mycelial growth (Guerriero et al., 2010; Rzeszutek et al. 2019a). Although NZ is an efficient inhibitor of CHSs in vitro, it has not been used in practice to control the pathogens due to the poor cellular uptake of the inhibitor by the hyphal cells.

a) b)

Fig. 17. Structures of nikkomycin Z (a) and UDP-N-acetylglucosamine (b).

Flupoxam, CGA and compound I

Flupoxam, CGA 325’615 and compound I (CI) are herbicides (Figure 18) (Hoffman and Vaughn, 1996; Vaughn and Turley, 2001; Peng et al., 2001; Sharples et al., 1998) that function as effective inhibitors of cellulose biosynthesis in plants. However, their precise mechanism(s) of action remain unknown. They reduce the incorporation of glucose into cellulose but at the same time increase its incorporation into different cell wall fractions like hemicelluloses or pectins, which suggests that glucose is diverted into other cell wall polysaccharides (García-Angulo et al., 2012). Moreover, the increased amount of glucose in polysaccharides tightly bound to cellulose most likely arises from soluble β-1,4-glucans (García- Angulo et al., 2012). This suggests that these herbicides inhibit crystallization of β-glucans rather than affecting glucose polymerization (García-Angulo et al., 2012). Flupoxam inhibits root elongation and causes changes in cell wall structure and composition (Hoffman and Vaughn, 1996; Vaughn and Turley, 2001). CI and CGA provoke a swollen phenotype in plant seedlings characteristic of cellulose synthesis inhibition (Kurek et al., 2002; Sharples et al., 1998). CGA prevents the synthesis of crystalline cellulose, and leads to rosette disruption and accumulation of non-crystalline β-1,4-glucans (Peng et al., 2001; Kurek et al., 2002). All three inhibitors affect the growth of S. parasitica. However, in vitro enzyme activity assays suggest that none of these inhibitors are specific for cellulose synthase (Rzeszutek et al., 2019c). This means that they probably do not bind to the catalytic subunit of the enzyme and inhibit cellulose biosynthesis in an indirect way. Moreover, stronger inhibitory effects on plant cell cultures than on the growth of S. parasitica mycelium indicates that these inhibitors are more specific to plants.

a) b)

Fig. 18. Structures of CGA (1-cyclohexyl-5-(2,3,4,5,6-pentafluoro- phenoxy)-1 λ4,2,4,6-thiatriazin-3-amine) (a), Compound I (5-tert-butyl- carbamoyloxyl-3-(3-trifluoromethyl) phenyl-4-thiazolidinone) (b) and Flupoxam (1-[4-chloro-3-[(2,2,3,3,3-pentafluoropropoxymethyl) phenyl]- 5-phenyl-1H-1,2,4-triazole-3-carboximide) (c).

DCB

DCB (2,6-dichlorobenzonitrile) (Figure 19) is a known inhibitor of cellulose biosynthesis. In plant cells, it inhibits the synthesis of cellulose at micromolar concentrations, leading to decreased amounts of cellulose in the cell wall (Meyer and Herth, 1978; Montezinos and Delmer, 1980; Mélida et al., 2009). The exact mode of action of this chemical is not fully understood, but several putative targets have been proposed. In Arabidopsis, DCB inhibits the mobility of cellulose synthase (CesA) complexes, which results in the accumulation of proteins at different regions of the plasma membrane (deBolt et al., 2007). A similar effect of increased amount of CesA proteins at the plasma membrane was also observed in tobacco cells (Nakagawa and Sakuraki, 1998). DCB was also shown to bind MAP20 in vitro. MAP20 is a microtubule-associated protein specifically up-regulated in secondary cell wall formation in plants (Rajangam et al., 2008). Moreover, a polypeptide of 18-kDa was identified as a possible receptor for DCB in plants and regulator of β- glucan synthesis (Delmer et al., 1987). DCB also affects cellulose biosynthesis in oomycetes. In P. infestans, DCB inhibits appressoria formation and prevents the infection of potato leaves, two processes that are dependent on cellulose synthesis (Grenville-Briggs et al., 2008). Additionally, DCB reduces the growth of S. monoica and provokes higher

34 expression levels of CesA genes in the mycelium as well as elevated glucan synthase activity in vitro (Fugelstad et al., 2009).

Fig. 19. Structure of 2,6-dichlorobenzonitrile (DCB).

Caspofungin

Caspofungin (Figure 20) belongs to the echinocandin family, a class of lipopeptide antifungal agents that act as non-competitive inhibitors of the GS complex. Caspofungin binds FKS1, the putative GS catalytic subunit in fungi. However, the exact mechanism of inhibition remains poorly understood (Douglas, 2001; Douglas et al., 1997; Munro, 2013). Disruption of β-1,3-glucan synthesis leads to the disintegration of the fungal cell wall and consequently to cell lysis. Caspofungin is highly effective against Candida and Aspergillus species and has been approved for clinical use (Denning, 2003). However, there are strains that have acquired mutations in the FKS gene and that show echinocandin resistance and higher chitin content in their cell walls. The treatment of cells with lower levels of caspofungin also leads to increased CHS activity and, consequently, increased chitin content in the cell wall, which results in lower susceptibility to the drug (Walker et al., 2008; Walker et al, 2013; Walker et al., 2015). Therefore, combining CHS inhibitors with caspofungin represents an interesting strategy to overcome the compensatory effect of CHS activity.

Fig. 20. Structure of caspofungin.

Biosurfactants as new promising inhibitors for the control of saprolegniosis

Massetolide A

Cyclic lipopeptides (CLP) are metabolites produced by various bacterial genera including Pseudomonas and Bacillus species and show activity against a broad range of microorganisms, including bacteria, fungi and oomycetes. The proposed mode of action of CLP is the formation of pores, leading to the disruption of membrane integrity and eventually cell death (Raaijmakers et al., 2010). Massetolide A is a CLP surfactant produced by the bacterial species Pseudomonas fluorescens (de Bruijn et al. 2008). It is composed of a cyclic oligopeptide consisting of 9 amino acids linked to the fatty acid tail of the molecule (Figure 21, Raaijmakers et al., 2006). Massetolide A shows activity against the human pathogenic bacteria Mycobacterium tuberculosis and Mycobacterium avium intercellulare but also shows strong activity against oomycetes (Gerard et al., 1997; van de Mortel et al., 2009). It causes lysis of the zoospores of the plant pathogenic oomycete P. infestans. Moreover, it also inhibits cyst germination, reduces mycelial growth and leads to increased hyphal branching and swelling (van de Mortel et al., 2009). Similar effects of massetolide A was observed in the case of S. parasitica. In the presence of this CLP, hyphal growth was inhibited and the cells exhibited increased branching and swelling (Liu et al., 2015). However, when tested in vivo,

36 massetolide A did not protect salmon eggs from colonization by Saprolegnia diclina (Liu et al., 2015).

Fig. 21. Structure of massetolide A.

The inhibitors mentioned above, especially the cell wall related inhibitors, represent powerful tools to control pathogens growth. Clinical use of caspofungin against fungal infections is one of the great examples of successful inhibition of cell wall formation. However, many of the inhibitors are not used in practice due to poor cellular uptake or unspecified mode of action. Therefore detailed knowledge about the mechanisms of cell wall biosynthesis and the identification of more specific and powerful inhibitors targeting cell wall biosynthetic enzymes are needed.

Aims of the thesis

The main objective of this work was to identify and characterize key enzymes involved in oomycete cell wall biosynthesis, to better understand the fundamental mechanisms of this process. Additionally, the impact of different inhibitors on the growth of S. parasitica, with an emphasis on cell wall related inhibitors, was investigated.

The specific aims of this research were to:

 Identify and characterize chitin and cellulose synthases from S. parasitica. Part of the work involved the use of cell wall related inhibitors such as nikkomycin Z, flupoxam, CGA and compound I (Papers I, II and IV).

 Express, purify and characterize SpCHS5, which has the highest expression level in the mycelium of S. parasitica (Paper III).

 Test bacterial biosurfactants as control agents of S. parasitica (Paper V).

Results and discussion

Paper I

Analysis of the genome of S. parasitica

S. parasitica is the first pathogenic oomycete of animals whose genome sequence was reported. The comparison of the generated sequence with already available genomes of pathogenic oomycetes of plants, such as Phytophthora and Pythium species, revealed distinctive gene expansions between the species and adaptation to specific hosts during evolution.

The genome size of S. parasitica was estimated to be 63 Mb. This is comparable with that of Py. ultimum (45 Mb), and P. ramorum (65 Mb), but significantly lower than the size of the genome of P. infestans (240 Mb). S. parasitica carries more than 17 000 coding genes, ~ 30 % of which exhibit loss of heterozygosity. Interestingly, the genome of S. parasitica possesses the highest gene density, the largest number of genes containing introns in comparison to other reported oomycetes, and one of the largest family of kinases reported to date.

Due to evolutionary adaptation to animal hosts, S. parasitica’s genome encodes a small number of cell wall degrading enzymes and lacks genes involved in inorganic nitrogen and sulfur assimilation.

In contrast, Saprolegnia contains a large number of secreted proteases, including serine, cysteine and metalloproteases, which represent potential virulence factors to attack host connective tissues and proteins. Indeed, one of the proteases (SPRG_14567) was shown to degrade fish immunoglobulin M (IgM), which suggests a suppressive function in immune responses for some of the proteases. Many of the proteases possess domain combinations specific to S. parasitica, for example peptidases with CBM1 (Carbohydrate Binding Module Family I) or ricin domains. It is possible that this specific domain architecture plays a role during pathogenesis. In addition to proteases, lectins represent the second largest group of secreted proteins. Lectins, as carbohydrate- binding proteins, most likely enhance the attachment of cysts to the host cells during infection stages through cell-cell interactions. Another

39 unique feature of S. parasitica’s genome is the presence of genes that code disintegrins, and galactose-binding lectins, which are typical for animal pathogens, but absent in oomycete pathogens of plants. The observed increased expression of these genes in cysts and germinating cysts indicate their potential role in pathogenesis. Moreover, expression analysis revealed that 10 % of all genes were induced in the pre-infection stages (cysts and germinating cysts) compared to the mycelium and sporulating mycelium. As opposed to Phytophthora species, S. parasitica lacks typical effector proteins, such as RXLR or Crinklers. The effector proteins enter plant cells where they promote infection mainly by suppressing plant immunity. S. parasitica contains a small group of host targeted proteins, one of which (SpHtp1) was shown to be translocated into the fish cells (van West et al., 2010). SpHtp1 does not share sequence similarity with any Phytophthora effectors.

Genome analysis also revealed the presence of six putative chitin synthase genes, whereas plant pathogenic oomycetes typically contain up to two Chs only. According to RNA-seq data, the expression of S. parasitica Chs genes is predominantly induced during the pre-infection phase, suggesting their specific role in pathogenesis.

Paper II

Characterization of chitin synthases from Saprolegnia parasitica

The analysis of the S. parasitica genome allowed the identification of six different chitin synthase genes, whose total number was confirmed by Southern blot analysis. The six chitin synthases, designated as CHS1 to CHS6, exhibited similar protein domain organization as the well known and catalytically active CHS2 from the yeast S. cerevisiae (Figure 22).

Fig. 22. Predicted domain organization of chitin synthase proteins from S. parasitica compared with CHS2 from the yeast S. cerevisiae.

All proteins contain multiple transmembrane domains at their C-terminal end, and a CHS domain carrying the conserved aspartic residues D, D, D and the pentapeptide QXXRW motif characteristic of most processive glycosyltransferases. Several oomycete CHS contain a MIT (Microtubule Interacting and Trafficking) domain, which is commonly present in many eukaryotic proteins, but identified for the first time in carbohydrate- active proteins (Guerriero et al., 2010). The investigation of the function of the MIT domain from the oomycete CHSs, suggests this domain is involved in the trafficking of the CHSs to the plasma membrane (Brown et al., 2016).

The occurrence of the specific chitin synthase motifs in oomycete CHSs points to potentially active enzymes. This was supported by in vitro activity assays, which demonstrated that the membrane proteins from S. parasitica’s mycelium produce chitin. The formation of the in vitro product was validated by hydrolysis with a specific chitinase enzyme.

Chitin synthase activity was detected in both microsomal fractions and detergent extracts prepared using either 0.5 or 1% CHAPS, dodecylmaltoside or digitonin. The enzyme activity in the detergent extracts, particularly in the presence of 1% digitonin, was highly enhanced in the presence of trypsin (~ 50 times), which acts as a proteolytic activator. As opposed to trypsin, nikkomycin Z strongly reduced chitin synthase activity in vitro. This is in agreement with the effect observed in vivo. The treatment with NZ resulted in a slower growth (Figure 23) and abnormal hyperbranched hyphae, which shows the importance of CHSs in the vegetative growth of S. parasitica.

Fig. 23. Growth inhibition of S. parasitica by 800 μM nikkomycin Z (NZ), a competitive inhibitor of chitin synthases.

Even though the mycelial growth was severely affected, it was not totally supressed. The disruption of chitin biosynthesis is possibly compensated by an up-regulation of some Chs genes to the level necessary for growth and viability. qPCR analysis revealed that four out of the six genes are expressed in the mycelium, i.e. Chs2, 3, 5 and 6, where Chs5 has the highest expression level, whereas Chs1 and Chs4 did not exhibit detectable expression. Only one of the four Chs expressed in the mycelium, Chs3, was up-regulated during NZ treatment. It is possible that CHS3 has an overlapping function with other S. parasitica CHSs involved in the vegetative growth and, most likely its up-regulation partially overcomes the effect of the inhibitor.

Paper III

Functional and biochemical characterization of SpCHS5

Chitin synthase 5 is one of the six different chitin synthases identified in S. parasitica. It exhibits the highest level of expression in the mycelium. This work describes for the first time the functional and biochemical characterization of an oomycete CHS through the successful expression and purification to homogeneity as a full-length protein containing all predicted transmembrane domains.

The gene coding for SpChs5 was cloned and transformed into the yeast S. cerevisiae, and successfully expressed as a fusion protein with the green fluorescence protein (GFP), in order to screen for protein expression, and a histidine tag (His-tag) for protein purification. The protein was purified using a nickel column, following detergent solubilization from the membrane. The expression and purity of the protein was analyzed using SDS-PAGE followed by in-gel fluorescence and Coomassie blue staining, which revealed the band corresponding to SpCHS5. The identity of the protein was verified by mass spectrometry analysis, with more than 50% of protein sequence coverage.

The catalytic activity of the recombinant enzyme was demonstrated using an in vitro radiometric assay. The results showed the presence of a radioactive product in the reaction mixture whose formation was highly inhibited by nikkomycin Z (Figure 24). The formation of chitin in vitro was demonstrated using a chitinase treatment, which resulted in complete degradation of the in vitro product. The characterization of chitin was further complemented by glycosidic linkage analysis, X-ray diffraction and electron microscopy, which altogether revealed the formation of chitin crystallites that consist of chains of chitin with an average length of 150 GlcNAc units (Figure 24).

The biochemical characterization of the enzyme revealed it is metal and pH dependent. The highest activity is observed in the presence of Mn2+ and at pH 7. The presence of metal ions is crucial for the enzyme. The ions stabilize and assist the departure of the leaving UDP group from the substrate. The activity of the enzyme was shown to be highly stimulated

43 by free GlcNAc and glycerol, which most likely act as allosteric activators. Moreover, the activity of the enzyme is higher after proteolytic treatment with trypsin, implying a zymogenic character of the protein.

Fig. 24. In vitro activity of the purified SpCHS5 showing inhibition with nikkomycin Z (NZ), and enzymatic hydrolysis of the chitin synthesized in vitro. The transmission electron micrograph shows the chitin crystallites formed in vitro.

Interestingly, our studies showed that SpCHS5 forms oligomers (Figure 25a). This was demonstrated using DMS (dimethyl suberimidate), a chemical crosslinker that reacts with primary amines. DMS connects only proteins that are in close proximity in a complex. After the crosslinking reaction, the main product observed was a dimer of CHS5 (~46% of the total protein). In addition, a band reflecting the presence of an oligomer of SpCHS5, most probably a trimer, was also visualized but with lower intensity (~30%). The quantification of the different forms of the proteins and the levels of activity retained after chemical crosslinking suggest that the dimer is the active form of the protein. However, other forms of the protein might also be active.

a) b)

Fig. 25. Oligomerization of CHS5 in the presence of the imidoester crosslinker DMS (in-gel fluorescence). A control reaction was prepared in the absence of DMS (a). High molecular mass protein complexes with an approximate size corresponding to the association of eight to ten SpCHS5 subunits detected by the Blue Native-PAGE (b).

In addition to chemical crosslinking, the use of native gel electrophoresis revealed the presence of oligomers (Figure 25b). In this case, the presence of larger protein complexes with the estimated size of eight to ten CHS5 subunits, was detected. The formation of oligomers has been observed for several glycosyltransferases. CHS oligomerization was previously observed in insects (Maue et al. 2009). Plant CesAs also oligomerize and the oligomers further assemble into higher molecular mass complexes (Doblin et al. 2002). The native gel shown in Figure 25b suggests that the oomycete CHS5 most likely assembles into larger complexes in a similar manner.

In order to further characterize the enzyme, the key amino acids involved in catalytic activity and processivity were identified by mutational analysis. The mutations of conserved residues in the active site completely abolished the activity of the enzyme, whereas the mutations of amino acids potentially involved in processivity (Con 2 region) provoked a reduction of the synthesis of insoluble chitin but the modified protein still retained the ability to form soluble chito-oligosaccharides (COS).

Paper IV

Characterization of the cellulose synthase genes in S. parasitica

The analysis of the genome sequence of S. parasitica revealed the presence of six putative cellulose synthase (CesA) genes. This number was confirmed by Southern blot analysis, which revealed the presence of three CesA2 (CesA2.1, CesA2.2 and CesA2.3), two CesA3 (CesA3.1 and CesA3.2) and one CesA4 genes. All S. parasitica CesA proteins were predicted to share common domain organization with plant and bacterial cellulose synthases (Figure 26), which suggest they are most likely active proteins.

Fig. 26. Predicted domain organization of CesA proteins from S. parasitica in comparison to bacterial (RsBcsA) and plant (AtCesA1) cellulose synthases.

The Saprolegnia proteins are characterized by the presence of a cellulose synthase domain flanked by multiple transmembrane regions and the

46 presence of the conserved motifs found in most processive glycosyltrasferases: D, D, D, and the pentapeptide QXXRW. Interestingly, the Saprolegnia CesA2 and CesA4 carry a plekstrin homology domain (PH) at their N-terminal ends. This domain is commonly present in many eukaryotic proteins, but the oomycete CesAs are the first carbohydrate- active enzymes where it was identified. The PH domain was functionally characterized in S. monoica’s CesA2 and shown to be involved in trafficking and/or targeting the enzyme to the plasma membrane (Fugelstad et al., 2012).

The expression of SpCesA genes was analysed by qPCR. Four of the six CesA genes, i.e. CesA2.2, CesA3.1, CesA4 and CesA2.1 are expressed in the mycelium, where CesA2.1 exhibits a significantly lower level of expression. The two other genes, CesA2.3 and CesA3.2 did not show any expression in the mycelium, which suggests that, together with CesA2.1, they are most likely involved during other stages of the life cycle. The highest expression level among all six SpCesA genes was observed for CesA3.1, whose orthologue was also shown to be highly expressed in other oomycetes, indicating an important role for this gene in cellulose formation.

The formation of cellulose was demonstrated using an in vitro radiometric assay using detergent-extracts of microsomal fractions from S. parasitica’s mycelium as a source of enzyme. The identity of the in vitro product was confirmed by glycosidic linkage analysis, which revealed the presence of 1,4-linked glucosyl residues.

To further characterize cellulose synthases, the effect of three inhibitors of cellulose biosynthesis, flupoxam, CGA 325’615 and compound I, was tested in vivo on the growth of the mycelium, as well as on the formation of cellulose in vitro and on gene expression. All three chemicals provoked a strong reduction of the growth of S. parasitica, by about 75% in the case of flupoxam and CGA 325’615, and 60% for compound I (Figure 27).

Fig. 27. Growth inhibition of S. parasitica by 200 μM of flupoxam, CGA and compound I.

The effect of the strongest inhibitors, flupoxam and CGA, was then tested on the expression of the genes that are potentially involved in cellulose biosynthesis. Only one gene, CesA3.1, was down-regulated in the presence of the drugs, the effect of which could be correlated to the growth inhibition, suggesting the involvement of this particular gene in cellulose biosynthesis in vivo. The tested inhibitors, however, were not found to be specific to CesA genes, but also affected the expression of Chs genes. Consistent with this observation, cellulose formation was inhibited together with chitin and β-1,3 glucan formation. This indicates that these inhibitors are not specific for cellulose synthase and most likely bind to other sites than the active site of the proteins, or indirectly interact with potential regulatory proteins that could be present in the detergent extracts.

Paper V

Activity of aquatic Pseudomonas species against Saprolegnia species

In this work, bacterial biosurfactants were tested as potential biological control agents against saprolegniosis.

Pseudomonas was found to be the largest group of bacteria associated with salmon eggs, both Saprolegnia-infected and healthy eggs. Around 80% of bacterial isolates from healthy eggs, and 60% from diseased eggs were able to inhibit the growth of S. parasitica and S. diclina in vitro. The

48 higher number of bacterial strains producing biosurfactants were isolated from healthy eggs.

When tested in vivo, seven out of eleven Pseudomonas isolates inhibited the infection of fish eggs by S. diclina. The highest activity against Saprolegnia was reported for the strain isolated from healthy eggs, called Pseudomonas strain H6. It was shown to produce a lipopeptide surfactant that is structurally similar to other biosurfactants, i.e. massetolide A and viscosin produced by Pseudomonas fluorescens SS101 and SBW25 strains, respectively.

The effect of this new, purified viscosin-like surfactant together with massetolide A was tested in vitro on the growth of S. parasitica and S. diclina. Both surfactants caused the formation of swollen and branched hyphae, but the compound produced by the H6 strain exhibited a stronger effect on the hyphal growth of Saprolegnia (Figure 28).

Fig. 28. Effect of different concentrations of biosurfactants (μg/ml) from Pseudomonas H6 and P. fluorescencs SS101 strains on the growth of S. diclina (a) and S. parasitica (b).

These purified compounds, however, when tested in vivo, did not protect salmon eggs from colonization by S. diclina. Nonetheless, these studies show the potential of using live organisms as biological control strategies to suppress the disease, where the biosurfactants likely contribute to the antimicrobial activity.

Conclusions and research perspectives

In this work, we identified and partially characterized chitin and cellulose synthases from S. parasitica, which sheds light on cell wall biosynthesis processes in oomycetes. Moreover, we demonstrated the relevance of cell wall synthesizing enzymes as targets for disease control.

Genome mining allowed us to identify several genes involved in cell wall biosynthesis, in particular the genes encoding chitin and cellulose synthases. We identified six chitin and six cellulose synthase genes. qPCR analysis revealed that four genes of each enzyme type were expressed in the mycelium, indicating their significance in vegetative growth. Indeed, the activity of CHSs and CesAs seems to be required for S. parasitica, since the treatment with cell wall related inhibitors, including NZ, flupoxam, CGA and CI, resulted in a strong growth inhibition. The analysis of RNA-seq data from different developmental stages of S. parasitica revealed that Chs genes were particularly highly induced during the pre-infection stage, indicating their specific role in pathogenesis. Thus, the analysis of chitin and cellulose synthases during different developmental stages can be very useful to understand their contribution and importance in the life cycle of the microorganism, hence in the infection process.

The presence of the characteristic conserved domains of glycosyltransferases was predicted in all CHS and CesA enzymes, indicating that they are potentially active proteins. We demonstrated that the proteins present in the mycelium were able to produce chitin and cellulose in vitro. However, in order to analyze if all the CHSs and CesAs genes in S. parasitica are coding for catalytically active proteins, a

50 detailed characterization of each individual enzyme would be necessary. This would also assist the development of additional specific inhibitors against CHSs and CesAs. In this work, we characterized one of the CHSs, namely SpCHS5, as a first oomycete chitin synthase. The protein was successfully expressed and purified to homogeneity as an active full-length protein. It was demonstrated that a single catalytic subunit, without any other CHS isoforms, is efficient to produce crystalline chitin in vitro. However, it seems to occur as a complex. Indeed, the recombinant CHS5 forms oligomers, mainly homodimers. Furthermore, our data suggest that CHS5 is able to assemble into larger multi-subunit complexes most likely forming terminal complexes in a similar manner as CesAs in plants. However, it cannot be concluded how many units are involved in the active complex and whether terminal complexes of the type found in plants (cellulose synthase rosettes) exist in oomycetes. The key amino acids essential for enzyme activity were identified by mutational analysis. This information could be very valuable in the development of new inhibition methods. This data would however need to be supported by the crystal structure of the protein to resolve the detailed mechanism of the enzyme. Taking into account the presence of multiple subunits in the CHS complex, this task is expected to be challenging.

In addition to cell wall inhibition, other interesting control strategies were presented in this work. These include the use of probiotic Pseudomonas bacteria and natural biosurfactants. Nonetheless, it has to be considered that some of the probiotic bacteria beneficial for one fish species might be inefficient or even harmful for others as well as for their eggs. Therefore, the influence of bacteria needs to be tested on various species of fish as well as on different fish life cycles.

Acknowledgements

It has been a long journey that finally has come to the end. Using this opportunity, I would like to express my gratitude to all without whom this work would have not been possible.

I am very grateful to Prof. Vincent Bulone for giving me the opportunity to work in the Glycoscience Division. I want to thank him for his supervision and guidance, helpful advice and all constructive discussions throughout all these years.

I give a special thanks to my co-supervisor Dr. Sara M. Díaz-Moreno for her invaluable help, support and patience with all my questions. I want also to thank all my colleagues, who contributed to the publications, especially Stefan and Osei.

Special thanks to Lauren for her valuable comments and critical editing of manuscripts.

Thank you Felicia for translating my abstract into Swedish.

I wish to thank all my present and former colleagues from the Glycoscience Division for their help, but also for creating a friendly and inspiring work atmosphere in the lab.

Dziękuję mojej rodzinie i przyjaciołom za wsparcie i wyrozumiałość w czasie moich studiów. Szczególne podziękowania dla mojej siostry Agnieszki za niocenioną pomoc i motywację podczas realizacji tej pracy.

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