Functional Characterization of Cellulose and Chitin Synthase Genes in Oomycetes

Functional characterization of cellulose and chitin synthase genes in Oomycetes

Johanna Fugelstad

Doctoral thesis in Biotechnology Division of Glycoscience Stockholm, Sweden 2011

School of Biotechnology Royal Institute of Technology AlbaNova University Center 106 91 Stockholm Sweden

Printed at US-AB universitetsservice

TRITA-BIO Report 2011:13 ISBN 978-91-7415-971-4 ISSN 1654-2312

Cover: Saprolegnia monoica mycelium stained with Congo Red (left) and human cell expressing the pleckstrin homology domain of S. monoica cellulose synthase 2, fused with green fluorescent protein (right). Micrographs by author. Johanna Fugelstad (2011) Functional characterization of cellulose and chitin synthase genes in Oomycetes. Doctoral thesis in Biotechnology. Royal Institute of Technology (KTH), Division of Glycoscience, Stockholm, Sweden. TRITA‐BIO Report 2011:13 ISBN 978‐91‐7415‐971‐4 ISSN 1654‐2312

ABSTRACT Some species of Oomycetes are well studied pathogens that cause considerable economical losses in the agriculture and aquaculture industries. Currently, there are no chemicals available that are environmentally friendly and at the same time efficient Oomycete inhibitors. The cell wall of Oomycetes consists of β‐(1Æ3) and β‐(1Æ6)‐glucans, cellulose and in some species minute amounts of chitin. The biosynthesis of cellulose and chitin in Oomycetes is poorly understood. However, cell wall synthesis represents a potential target for new Oomycete inhibitors. In this work, cellulose and chitin synthase genes and gene products were analyzed in the plant pathogen Phytophthora infestans and in the fish pathogen Saprolegnia monoica.

A new Oomycete CesA gene family was identified, containing four subclasses of genes designated as CesA1 to 4. The gene products of CesA1, 2 and 4 contain pleckstrin homology (PH) domains located at the N‐terminus, which is unique to the Oomycete CesAs. Our results show that the SmCesA2 PH domain binds to phosphoinositides, F‐actin and microtubules in vitro and can co‐localize with F‐actin in vivo. Functional characterization of the CesA genes by gene silencing in P. infestans led to decreased cellulose content in the cell wall. The cellulose synthase inhibitors DCB and Congo Red inhibited the growth of the mycelium of S. monoica and had an up‐regulating effect on SmCesA gene expression. Zoospores from P. infestans treated with DCB were unable to infect potato leaves. In addition, two full‐length chitin synthase genes (Chs) were analyzed from S. monoica. Expression of SmChs2 in yeast yielded an active recombinant protein. The biochemical characterization of the in vitro product of SmChs2 confirmed that the protein is responsible for chitin formation. The chitin synthase inhibitor nikkomycin Z inhibited the SmChs2 both in vivo and in vitro.

Altogether these results show that at least some of the CesA1‐4 genes are involved in cellulose biosynthesis and that synthesis of cellulose is crucial for infection of potato by P. infestans. The PH domain is involved in the interaction of CesA with the cytoskeleton. In addition, we firmly demonstrate that the SmChs2 gene encodes a catalytically active chitin synthase.

Keywords: cellulose biosynthesis; chitin biosynthesis; cellulose synthase genes; chitin synthase genes; Oomycetes; Phytophthora infestans; Saprolegnia monoica; pleckstrin homology domain.

© Johanna Fugelstad, 2011 Johanna Fugelstad (2011) Funktionell karaktärisering av cellulosa‐ och kitinsyntasgener i oomyceter. Doktorsavhandling i Bioteknologi. Kungliga Tekniska högskolan, avdelningen för Glykovetenskap, Stockholm, Sverige. TRITA‐BIO Report 2011:13 ISBN 978‐91‐7415‐971‐4 ISSN 1654‐2312

SAMMANFATTNING Många oomyceter är välstuderade patogena arter som orsakar ansenliga ekonomiska förluster inom jordbruks‐ och fiskodlingsindustrin. För närvarande finns inga kemikalier till hands som effektivt bekämpar oomyceter och på samma gång är miljövänliga. Oomyceters cellvägg består av β‐(1Æ3)‐ och β‐(1Æ6)‐glukaner, cellulosa och i vissa arter små mängder kitin. Hur oomyceter syntetiserar cellulosa och kitin är relativt okänt, dock är cellväggssyntesen ett potentiellt mål för nya oomycetinhibitorer. Cellulosa‐ och kitinsyntasgener och dess genprodukter, från växtpatogenen Phytophthora infestans och fiskpatogenen Saprolegnia monoica, har analyserats i den här avhandlingen.

En ny familj cellulosasyntasgener från oomyceter har identifierats, innehållande fyra underklasser av gener, utsedda till CesA1 till 4. N‐terminalerna i genprodukterna från CesA1, 2 och 4 innehåller en pleckstrinhomolog(PH)‐domän, vilket är unikt för CesAs från oomyceter. Våra resultat visar att PH‐domänen från SmCesA2 binder till fosfoinositider, F‐aktin och mirotubuli in vitro och kan samlokalisera med F‐aktin in vivo. Tystande av CesA‐generna i P. infestans ledde till en minskad mängd cellulosa i cellväggen. Utöver detta inhiberade cellulosasyntasinhibitorerna DCB och Kongo Rött tillväxten av S. monoica mycel och hade en uppreglerande effekt på uttrycket av SmCesA‐generna. Zoosporer från P. infestans behandlade med DCB var oförmögna att infektera potatisblad. Dessutom analyserades två fullängds kitinsyntasgener (Chs) från S. monoica. Rekombinant uttryck av SmChs2 i jäst resulterade i ett aktivt protein. Den biokemiska analysen av in vitro‐produkten från SmChs2 bekräftade att proteinet syntetiserar kitin. Kitinsyntasinhibitorn nikkomycin Z inhiberade SmChs2 både in vivo och in vitro.

Sammataget visar dessa resultat att åtminstone några av generna CesA1‐4 är involverade i cellulosabiosyntes och att syntes av cellulosa är nödvändigt för P. infestans infektionsförmåga. PH‐domänen är involverad i interaktionen mellan CesA och cytoskelettet. Dessutom har vi bevisat att SmChs2‐genen kodar för ett katalytiskt aktivt kitinsyntas.

Nyckelord: cellulosabiosyntes; kitinbiosyntes; cellulosasyntasgen; kitinsyntasgen, Oomyceter, Phytophthora infestans; Saprolegnia monoica; pleckstrinhomolog‐domän

LIST OF PUBLICATIONS

Publication I Grenville‐Briggs L. J., Anderson V. L., Fugelstad J., Avrova A. O., Bouzenzana J., Williams A., Wawra S., Whisson S. C., Birch P. R., Bulone V. and van West P., Cellulose synthesis in Phytophthora infestans is required for normal appressorium formation and successful infection of potato, 2008, Plant Cell 20(3):720‐38

Publication II Fugelstad J., Bouzenzana J., Djerbi S., Guerriero G., Ezcurra I., Teeri T.T., Arvestad L. and Bulone V. Identification of the cellulose synthase genes from the Oomycete Saprolegnia monoica and effect of cellulose synthesis inhibitors on gene expression and enzyme activity, 2009, Fungal Genet. Biol. 46(10):759‐67

Publication III Fugelstad J.*, Brown C.*, Hukasova E., Sundqvist G., Lindqvist A. and Bulone V., Functional characterization of the pleckstrin homology domain of a cellulose synthase from the Oomycete Saprolegnia monoica, 2011, Manuscript

Publication IV Guerriero G., Avino M., Zhou Q., Fugelstad J., Clergeot P.H. and Bulone V. Chitin synthases from Saprolegnia are involved in tip growth and represent a potential target for anti‐oomycete drugs, 2010, PLoS Pathog. 6(8): 1‐12

*Both authors contributed equally to the work

AUTHOR’S CONTRIBUTION

Publication I: Johanna Fugelstad’s contribution is the biochemical characterization of the cell wall, participation in the co‐localization studies using antibodies and some of the writing.

Publication II: Johanna Fugelstad did most of the experimental work and the writing with assistance from Dr. Lars Arvestad for the bioinformatic sections.

Publication III: Johanna Fugelstad performed the cloning, expression, purification, bioinformatics and parts of the localization of the experimental section, and wrote the manuscript with assistance from Christian Brown in the Materials and Methods section.

Publication IV: Johanna Fugelstad’s contribution is the Southern blot experiment and participation in some of the writing.

RELATED PUBLICATIONS NOT INCLUDED IN THE THESIS

Haas B.J., Kamoun S., Zody M.C., Jiang R.H., Handsaker R.E., Cano L.M., Grabherr M., Kodira C.D., Raffaele S., Torto‐Alalibo T., Bozkurt T.O., Ah‐Fong A.M., Alvarado L., Anderson V.L., Armstrong M.R., Avrova A., Baxter L., Beynon J., Boevink P.C., Bollmann S.R., Bos J.I., Bulone V., Cai G., Cakir C., Carrington J.C., Chawner M., Conti L., Costanzo S., Ewan R., Fahlgren N., Fischbach M.A., Fugelstad J., Gilroy E.M., Gnerre S., Green P.J., Grenville‐Briggs L.J., Griffith J., Grünwald N.J., Horn K., Horner N.R., Hu C.H., Huitema E., Jeong D.H., Jones A.M., Jones J.D., Jones R.W., Karlsson E.K., Kunjeti S.G., Lamour K., Liu Z., Ma L., Maclean D., Chibucos M.C., McDonald H., McWalters J., Meijer H.J., Morgan W., Morris P.F., Munro C.A., O'Neill K., Ospina‐Giraldo M., Pinzón A., Pritchard L., Ramsahoye B., Ren Q., Restrepo S., Roy S., Sadanandom A., Savidor A., Schornack S., Schwartz D.C., Schumann U.D., Schwessinger B., Seyer L., Sharpe T., Silvar C., Song J., Studholme D.J., Sykes S., Thines M., van de Vondervoort P.J., Phuntumart V., Wawra S., Weide R., Win J., Young C., Zhou S., Fry W., Meyers B.C., van West P., Ristaino J., Govers F., Birch P.R., Whisson S.C., Judelson H.S., Nusbaum C., Genome sequence and analysis of the Irish potato famine pathogen Phytophthora infestans, 2009, Nature, 461(7262):393‐8.

Guerriero G., Fugelstad J. and Bulone V., What do we really know about cellulose biosynthesis in higher plants? 2010, J. Integr. Plant Biol. 52(2):161‐75, review

LIST OF CONTENTS

1 INTRODUCTION 1 2 INTRODUCTION TO OOMYCETES 1 2.1 General characteristics of Oomycetes 1 2.2 Plant pathogens of the genus Phytophthora 3 2.3 Fish pathogens of the genus Saprolegnia 6 2.4 Genomic data on Oomycete species 7 2.5 Multidomain proteins of Phytophthora species 8 3 CELL WALLS 9 3.1 Oomycete cell walls 9 3.2 Plant cell walls 10 3.3 Fungal and yeast cell walls 11 4 INTRODUCTION OF CELLULOSE AND CHITIN 13 4.1 Cellulose, an important carbohydrate 13 4.2 Chitin 16 5 CELLULOSE AND CHITIN BIOSYNTHESIS IN DIFFERENT ORGANISMS 18 5.1 Catalytic subunits of cellulose and chitin synthases 18 5.1.1 Classification and properties of cellulose and chitin synthases 18 5.1.2 Inverting mechanism of GT2s 20 5.1.3 Pleckstrin homology domains 22 5.2 Cellulose biosynthesis 26 5.2.1 Terminal complexes responsible for cellulose biosynthesis 26 5.2.2 Cellulose biosynthesis in bacteria 27 5.2.3 Cellulose biosynthesis in higher plants 28 5.2.4 Cellulose biosynthesis in Oomycetes 37 5.2.5 Cellulose biosynthesis in other organisms 38 5.2.6 Biochemical approaches to study cellulose synthesis in vitro 38 5.3 Chitin biosynthesis 41 5.4 Inhibitors of glycosyltransferase activities 45

6 PRESENT INVESTIGATION 48 6.1 Aim of the present investigation 48 6.2 Materials and methods 49 6.3 Results and discussion 50 6.3.1 Characterization of a novel group of CesA genes in Oomycetes (Papers I, II and unpublished results) 50 6.3.2 Expression pattern, localization and function of the CesAs from P. infestans (Paper I) 54 6.3.3 Effect of cellulose synthesis inhibitors on mycelial growth of S. monoica, expression of CesA genes and glucan synthase activities (Paper II) 58 6.3.4 Characterization of the PH domain from SmCesA2 (Paper III) 62 6.3.5 Characterization of the Chs genes in the Oomycetes S. monoica and S. parasitica (Paper IV and unpublished material) 64 6.3.6 In vitro chitin synthase activities of recombinant SmChs1 and SmChs2 (Paper IV) 65 6.4 Conclusions and perspectives 67 7 ACKNOWLEDGEMENTS 70 8 REFERENCES 72

1. INTRODUCTION This thesis is based on work on several organisms and spanning a broad spectrum of techniques. The common thread is two of the most important carbohydrates on earth, namely cellulose and chitin. The aim of the present investigation is to understand the mechanism and function of cellulose and chitin biosynthesis in Oomycetes, by investigating the corresponding synthesizing enzymes and a subset of their functional domains. To guide the reader to the work presented in the thesis, the introduction starts by describing the basic biology of Oomycetes. It then continues by discussing the function and properties of the plant, fungal and Oomycete cell walls. In the following part, cellulose and chitin are presented in more detail, especially their function, structure and biosynthesis. In the present work, some functional domains of Oomycete cellulose synthesizing enzymes have been investigated. Therefore, some background necessary to understand the characteristics of these domains is presented. Since the work is focused on cellulose and chitin biosynthesis, the final part of the introduction reviews the subject and gives an insight on the current knowledge in the field.

2. INTRODUCTION TO OOMYCETES

2.1 General characteristics of Oomycetes Oomycetes were previously classified in the kingdom Fungi, but in the last decades it has been shown that they in fact belong to the Stramenopiles and are related to heterokont algae, such as for instance brown algae (Kumar and Rzhetsky, 1996; Paquin et al., 1997; Baldauf et al., 2000). The growth of Oomycetes resembles the growth of fungal cells that form coenocytic hyphae. The cell wall of Oomycetes, as opposed to that of fungi, contains cellulose. Most stages in the Oomycete life cycles are diploid (Hardham, 2007). Oomycetes have evolved a saphrophytic and sometimes pathogenic life style. Some species combine both life styles, and have adapted to many types of host organisms such as for instance plants, nematodes, vertebrates and crustaceans (Table 1). Oomycetes can be biotrophs, hemibiotrophs or necrotrophs. Biotrophic species obtain their nutrients from living tissues while hemibiotrophs first feed on living tissues and kill their host at a later stage.

Necrotrophs live exlusively on dead host tissues. The plant pathogenic Oomycetes have been most characterized while much less is known about the animal pathogens. Phytophthora infestans, the cause of potato blight, belongs to the well studied plant pathogenic genus Phytophthora (Agrios, 2005). More than 100 years after the infamous Irish famine caused by the potato late blight, Oomycetes still represent an important problem in agriculture. The losses due to plant pathogenic Phytophthora species have constantly increased since the dissemination of the second mating type that lead to genetic recombinations and the emergence of more virulent strains of P. infestans (Fry and Goodwin, 1997). The spread of Oomycetes to new habitats has lead to new diseases and devastation of some crops. One example is Phytophthora ramorum, the agent of Sudden Oak Death, killing coastal oaks in California (Rizzo et al., 2002). There is currently no efficient and environmentally friendly drug that prevents or stops infections by plant pathogenic Oomycetes. Not even anti‐fungal drugs allow the control of the diseases caused by Oomycetes (Hardham, 2007). The growing business of aquaculture is seriously affected by Oomycetes, particularly by different species of the order Saprolegniales. Outbreaks of Saprolegniosis are responsible for losses of up to 50 % (Meyer, 1991). Malachite green, used until recently to control Saprolegnia, was banned worldwide in 2002 because of its carcinogenic and toxicological effect. Currently, there are no drugs available that are at the same time efficient and safe for the fish host and for human consumption of treated fish (van West, 2006). P. infestans and Saprolegnia monoica were chosen as representative model organisms for the work presented here due to their economical importance and impact as plant and animal pathogens, respectively. However, the economical and biological importance of other Oomycetes should be stressed, although they are not included in the scope of this study.

Table 1. Examples of pathogenic species from different orders of Oomycetes, their specific hosts and corresponding diseases. The phylogenetic relationship between Oomycetes shows that animal pathogens are found in several genera, except the Peronosporales, which contains strictly plant pathogens (Phillips et al., 2008).

Orders and representative species Hosts and diseases of different genera Saprolegniales Saprolegnia monoica Saprolegniosis in fish Saprolegnia parasitica Saprolegniosis in fish Achlya bisexualis Ulcerative mycosis in fish Aphanomyces astaci Plague in crayfish Aphanomyces euteiches Root rot disease in pea

Lagenidiales Lagenidium giganteum Infects mosquito larvae

Pythiales Pythium insidiosum Skin lesions in mammals Pythium oligandrum Infects fungi

Peronosporales Hyaloperonospora parasitica Downy mildew in plants Phytophthora infestans Late blight in potato Phytophthora sojae Stem and root rot in soybean Albugo candida White rust in plants

2.2 Plant pathogens of the genus Phytophthora The plant pathogenic genus Phytophthora includes hemibiotrophs (e.g. P. infestans, P. sojae) and necrotrophs (e.g. P. cinnamomi). Phytophthora species have had a severe impact on agriculture and have served as model organisms to study plant pathology (Hardham, 2007). 3

A well known and typical example is P. infestans (Figure 1). This microorganism can infect a variety of plant species in addition to potato, such as tomato and sojae. It probably has its origin in Mexico (Andrivon, 1996) and followed the spread of potato all over the world (Schumann, 1991). P. infestans can infect the whole potato plant, tubers, roots or leaves, leading to the death of the plant.

Figure 1. Late blight on a potato leaf caused by P. infestans. Photo from paper I (reprinted with permission from Grenville‐Briggs et al., 2008).

Life cycle and reproduction of Phytophthora species The life cycle of Oomycetes includes sexual and asexual reproduction, with flagellated free swimming spores characteristic of Stramenopiles (Figure 2 A). The mycelium of P. infestans produces branched sporangiophores containing lemon‐shaped asexual sporangia at their tips. When the sporangium bursts it releases motile zoospores with two flagella, which is typical for heterokonts. The role of the zoospores is transmission of the pathogen from host to host. They are also essential for targeting the site of infection (Walker and van West, 2007). The host attracts the zoospores by chemotactic and electrotactic stimuli (Tyler, 2002). Spores can also be transported to adjacent places by the wind, explaining their fast dissemination. However, the spores need humid conditions to be able to infect their hosts

(Hardham, 2007; Walker and van West, 2007). The zoospores lack cell walls and maintain their volume and homeostasis with the help of a contractile vacuole (Mitchell and Hardham, 1999). Upon contact with a host, the zoospores become immotile and encyst, which leads to the formation of a cell wall. A germ tube protrudes from the cyst forming an appressorium that resembles a swollen tip of the germ tube. The appressorium penetrates the host tissue directly or enters via stomata in the leaf (Figure 2 B), and forms an infection vesicle. The mycelium then grows from the infection vesicle in between the plant cells, sometimes with haustoria, which is a mycelium that has penetrated the host cell wall and invaginated the plasma membrane (Grenville‐Briggs and van West, 2005). Older infected plant cells die while the mycelium continues to spread into the host tissue (Agrios, 2005; Hardham, 2007). The sexual reproduction of Oomycetes results in thick walled resistant oospores that can survive for 3‐4 years outside a host (Figure 2 A). Oospores are formed when the male reproductive organ, the antheridium, fertilizes the female reproductive organ, the oogonium. For oospore formation in P. infestans to take place, strains of two mating types are needed, A1 and A2. It was not until recently that the A2 mating type emerged from Mexico and spread in the rest of the world, thereby making possible the sexual reproduction of Phytophthora. This has lead to the emergence of more virulent strains due to genetic recombinations of pathogenic characteristics of the mating strains (Agrios, 2005).

A B

Figure 2. A) Life cycle of P. infestans with sexual and asexual sporulation (adapted from Schumann, 1991). B) Germinated cyst of P. infestans with its appressorium on a potato leaf and a stomata in the bottom left corner (from paper I, reprinted with permission from Grenville‐Briggs et al., 2008).

2.3 Fish pathogens of the genus Saprolegnia Members of the genus Saprolegnia cause the group of diseases called Saprolegniosis on fish or fish eggs. Saprolegniosis are characterized by visible white or gray patches of filamentous mycelium on the fish (Lartseva, 1986). The infection starts by the invasion of the epidermal tissue and can spread over the entire body (Figure 3 A). Saprolegnia species are considered as opportunistic pathogens, which have a saprophytic and necrotrophic life style (Neish, 1977; van West, 2006). There is currently no efficient treatment of Saprolegniosis available. Good fish management during aquaculture can decrease the risk of infection since stress seems to be an important risk factor for Saprolegniosis (Willoughby and Pickering, 1977; Meyer, 1991).

Life cycle and reproduction of Saprolegnia species The life cycle of Saprolegnia species is similar to that of other Oomycetes, with the ability to produce asexual and sexual spores. The asexual reproduction of Saprolegnia includes a special feature called polyplanetism. The asexual spores release primary motile zoospores that within a short time encyst and release secondary zoospores (Figure 3 B). The liberation of secondary zoospores favours multiple attempts to locate a suitable host. In particular, these spores are motile for a longer period than primary zoospores and possess hairs, also called boat hooks, which are believed to increase attachment efficiency. These hairs are unique to the genus Saprolegnia (van West, 2006).

A B

Figure 3. A) Wild salmon infected by S. parasitica with white infected lesions on the skin. B) Life cycle of S. parasitica (pictures adapted from van West, 2006).

2.4 Genomic data on Oomycete species More knowledge on the molecular events occurring during the life cycle of Oomycetes is needed to develop tools that allow the control of their dissemination and infection

7 processes. The genomes of three economically important plant pathogens, namely Phytophthora infestans, Phytophthora sojae and Phytophthora ramorum have been fully sequenced at the Broad Institute (http://www.broad.mit.edu/; Tyler et al., 2006a) and the Joint Genome Institute (http://www.jgi.doe.gov/), opening new opportunities to characterize these virulent species (Tyler et al., 2006b). An EST database is also available for several species, including members of the Peronosporales and Saprolegniales such as S. parasitica and P. infestans (www.oomycete.org; Gajendran et al., 2006). The first draft of the S. parasitica genome was made publically available in 2009 (www.broadinstitute.org). The genome of the Oomycete Hyaloperonospora arabidopsidis is also sequenced (Baxter et al., 2010). An EST library with more than 1510 ESTs sequenced from a mycelial cDNA library of S. parasitica is available (Torto‐Alalibo et al., 2005). The average amino acid identity between S. parasitica and the three Phytophthora species whose genomes have been sequenced, based on the alignment of 18 deduced sequences of conserved proteins, is of 77 % compared to 93 % among the Phytophthora species (Torto‐Alalibo et al., 2005).

2.5 Multidomain proteins of Phytophthora species The genomes of P. sojae and P. ramorum contain a significant amount of novel multifunctional proteins, with combinations of domains not present in bacterial, plant, fungal or animal genomes (Morris et al., 2009). Many of the fusion proteins are active in primary metabolic or regulatory pathways. A large proportion of the novel combinatorial proteins contain a calcium‐binding domain, a protein‐binding domain, a zinc finger domain or a kinase related domain (Morris et al., 2009). The presence of new combinations of domains in kinases was also found when the P. infestans genome was investigated with bioinformatic tools (Judelson and Ah‐Fong, 2010). The phylogenetic origin of multifunctional protein fusions is not clear, but it is believed that bacterial horizontal gene transfer as well as genes from a photosynthetic endosymbiont contributed. Gene transfer occurred before the split of the diatom and Oomycete lineages giving rise to this unique feature of Stramenopile genomes (Morris et al., 2009). The multidomain proteins are strictly conserved between P. sojae and P. ramorum, further supporting this theory, although not every protein class is characterized by an increase in the number of multidomain proteins. Among the P. infestans kinases, the amount of multidomain proteins is decreased compared to the human genome

(Judelson and Ah‐Fong, 2010). This is believed to be related to a higher degree of transcriptional regulation of kinases in P. infestans compared to post‐translational regulation in humans. Also, the protein kinase B family in P. infestans contains fewer proteins with pleckstrin homology domains compared to humans. Rather than the number of modular proteins, it is the number of novel combinations of domains that is believed to be unique to Stramenopiles, especially Oomycetes.

3. CELL WALLS

3.1 Oomycete cell walls The cell wall of Oomycetes functions as a barrier for the cell against the surrounding milieu. It determines the cell shape, adapts with the life cycle and is important for the infection of pathogenic Oomycetes. The cell walls of some Oomycetes (e.g. S. monoica and species of Apodachlya) contain chitin, like fungal cell walls (see section on Fungal and yeast cell walls, paragraph 3.3), but in a much smaller proportion, representing less than 1 % of the total cell wall carbohydrates as opposed to up to ~20 % of the dry weight in some fungi (Bartnicki‐ Garcia, 1968; Lin and Aronson, 1970; Aronson and Lin, 1978; Bulone et al., 1992). The major cell wall polysaccharides in Oomycetes are β‐(13)‐glucans and β‐(16)‐glucans as well as cellulose, which has never been reported in any fungal species (Bartnicki‐Garcia, 1968). The Oomycete cellulose is microfibrillar, of low crystallinity, and corresponds to cellulose IV (Bulone et al., 1992; Helbert et al., 1997), which is a disorganised form of cellulose I. The mode of assembly of the carbohydrate components in the cell walls of Oomycetes is not known, although it is believed that the cell walls consist of an inner layer of cellulose microfibrils doubled by an amorphous matrix composed essentially of other β‐glucans (Bartnicki‐Garcia, 1968). Cellulose in Oomycetes has most likely the same scaffolding function as chitin in fungal cell walls. The chitin in the cell walls of some Oomycetes is of the α type, as shown by electron diffraction analysis, and occurs as granular structures in the cell wall as well as after chemical extraction (Bulone et al., 1992). In some Oomycete species, although the cell wall contains GlcNAc‐based carbohydrates, there is little evidence of the occurrence of crystalline chitin (Werner et al., 2002). For instance, in Aphanomyces euteiches about 10 % of the cell wall consists of GlcNAc‐based carbohydrates, which correspond to

9 soluble chitosaccharides rather than crystalline chitin (Badreddine et al., 2008). Oomycetes belonging to the Phytophthora genus are devoid of chitin.

3.2 Plant cell walls A general description of the plant cell walls is briefly presented here to facilitate the understanding of the comparisons made in the thesis. Cell walls are divided into two major categories, type I and type II, based on their structures and components. This is a broad generalisation since the cell wall composition fluctuates considerably among different plant species. In type I cell walls characteristic for dicots (Figure 4), the major hemicellulose is xyloglucan, whereas in type II cell walls the major hemicelluloses are mixed linked glucans (β‐(13,14)‐glucans) and glucuronoarabinoxylans. Type II cell walls are characteristic of the Poales family, which comprises grasses and cereals. It should however be noted that the cell walls differ even throughout one individual plant depending on tissue and cell type and even within one cell depending on growth and differentiation stage (Keegstra, 2010). Plant cell walls consist of a complex matrix of cellulose microfibrils, hemicelluloses, pectins and structural proteins (Figure 4) (for a review on plant cell walls, see for instance Cosgrove, 2005). There are three major roles for the plant cell wall: to support the cell shape, to protect the protoplasm and act as a first defence for the plant cell. The structure of the cell, which depends on cell type, is determined by the cell wall. The cell wall protects the cell from dehydration and ultra‐violet radiation. The cell wall is the first line of defence against microbial attacks. Some cells, especially in vascular tissue in woody plants, contain a thick secondary cell wall enriched in cellulose and lignin.

Figure 4. Model of a type I primary plant cell wall (after Carpita and Gibeaut, 1993). In type II cell walls, the major hemicelluloses are mixed linked glucans (β‐(13,14)‐glucans) and glucuronoarabinoxylans instead of xyloglucan.

3.3 Fungal and yeast cell walls A general description of the fungal and yeast cell walls is briefly presented here to facilitate the understanding of the comparisons made in the thesis. Fungi and yeasts belong to the Fungi kingdom and their cell walls consist mostly of an ordered mesh of branched β‐(13)‐ and β‐(16)‐glucans, chitin, mannans and mannoproteins (Figure 5) (Bartnicki‐Garcia, 1968). The cell wall is essential for the life of the fungi. The glucans comprise the major part of the cell wall dry weight in yeast (Bartnicki‐Garcia, 1968). The β‐(13)‐glucan chains are branched with β‐(16)‐linkages, as opposed to the linear callose in plants. Since β‐(13)‐ glucans are the major components of the cell wall, their synthesis is vital for the yeast. The

11 catalytic subunit of the yeast β‐(13)‐glucan synthase is encoded by the FK506‐ hypersensitive locus (FKS) (Douglas et al., 1994). In Saccharomyces cerevisiae there are two copies, FKS1 and FKS2 (Douglas et al., 1994; Mazur et al., 1995). The two FKS isoforms are differently regulated. FKS1 is mostly expressed when cells are grown with glucose as the carbon source, whereas FKS2 is expressed when the medium lacks glucose. Calcium triggers the expression of FKS2. Double knockouts of FKS1 and FKS2 are lethal and the proteins are partially redundant. FKS1 and FKS2 have different roles in the cell cycle of yeast. FKS2 is important for sporulation while FKS1 is the most expressed isoform during vegetative growth (Mazur et al., 1995). FKS activity is regulated by Rho1p, a GTPase. Rho1p is needed for the full activity of the FKS complex (Kang and Cabib, 1986; Qadota et al., 1996). The proteins in the cell wall are mainly glycoproteins. In the yeast, these are O‐ and N‐ glycosylated mannoproteins. Most of the proteins are covalently bound via glycosidic linkages to the chitin or glucans present in the cell wall (Bowman and Free, 2006). β‐(16)‐ Glucans are structurally important as they crosslink the mannoproteins to β‐(13)‐glucans and chitin (van der Vaart et al., 1996). To date, little is known about the synthesis of β‐ (16)‐glucans and there is no catalytic subunit identified for this activity. Several proteins, like the killer toxin resistant (KRE) proteins, have been linked to β‐(16)‐glucan synthesis. A deletion in KRE5 leads to a complete lack of β‐(16)‐glucans (Shahinian and Bussey, 2000). The protein shows similarities to a UDP‐glucose:glycoprotein glucosyltransferase and is localized in the endoplasmatic reticulum (ER), but the biochemical activity of KRE5 is unknown (Shahinian and Bussey, 2000). Deletion in other KRE proteins lead to decreased β‐ (16)‐glucan levels (up to 50 % decrease), but the precise role of the proteins in β‐(16)‐ glucan biosynthesis is unclear. Chitin constitutes only a minor part of yeast cell walls, about 1‐2 % of the dry weight. S. cerevisiae and Pichia pastoris have three chitin synthase genes, Chs1, Chs2 and Chs3 (Roncero, 2002; De Schutter et al., 2009). These genes have been characterized by mutant complementation in S. cerevisiae and their function will be discussed in Function and regulation of yeast Chs from classes I, II and IV in paragraph 5.3.

Figure 5. Model of a yeast cell wall. The major constituents of the yeast cell wall are β‐(13)‐glucans, β‐(16)‐glucans, chitin and mannoproteins, which are sometimes attached to the membrane by a glycosylphosphatidylinositol (GPI)‐anchor.

4. INTRODUCTION TO CELLULOSE AND CHITIN In this part, basic knowledge on the structure, function and natural sources of cellulose and chitin is presented, as well as the main industrial applications of the carbohydrates.

4.1 Cellulose, an important carbohydrate Cellulose has been used as a chemical raw material for about 150 years and, historically, it has been utilized by humans since the time of pharaohs in the form of papyrus. Half of the total annual biomass produced by algae, plants and some bacteria is composed of cellulose, making it the most abundant macromolecule on earth and pointing out the biological importance of the polymer. Cellulose is the building material of the cell walls of plants, but it also occurs in other organisms such as algae and tunicates, providing mechanical support and protection. In plants, cellulose is deposited in a primary cell wall together with hemicelluloses, pectins and structural proteins. Woody tissues contain a thick secondary cell

13 wall made of crystalline cellulose. The biological importance of cellulose for bacteria and other organisms are discussed later. Cellulose is currently considered as an alternative source of energy and chemicals to fossil fuels. Cellulose also represents an abundant raw material that can potentially satisfy the increasing demand for environmentally friendly and biocompatible products, and it is already used in fabrics and paper. The biosynthesis of cellulose has been exploited for the development of new biomaterials. One typical example is the use of bacterial cellulose for the formation of artificial blood vessels and wound dressings (Klemm et al., 2005; Bodin et al., 2007). However, plants remain the main source of cellulose. For instance, the cotton fiber produces virtually pure cellulose and is largely utilized in the textile industry. Cellulose and its derivatives, e.g. nitrocellulose, cellulose acetate, methyl cellulose, carboxymethyl cellulose, have numerous industrial applications, for instance in the form of coatings, pharmaceuticals, cosmetics and adhesives (Klemm et al., 2005).

Cellulose structure Cellulose is a linear homopolymer of D‐glucose residues linked through β‐(14) bonds. The 4 D‐glucose residues are in the C1 chair conformation. The cellulose molecule has a chemical polarity: one end of the β‐glucan chain carries a free hemiacetal group with reducing properties and is referred to as the reducing end. The anomeric carbon of the D‐ glucopyranose unit at the other end is involved in a glycosidic bond with the penultimate residue. This end of the chain is referred to as the “non‐reducing” end (Figure 6). Since the cellulose molecule consists of β‐(14) linkages, each of the glucose moieties are inverted almost 180° relative to their adjacent moieties. For this reason, cellobiose is sometimes considered as the repeating unit of cellulose as represented in Figure 6. The degree of polymerization (DP) of cellulose varies depending on the origin of the polymer and the extraction protocol. In wood pulp, the DP ranges typically between 300 and 1700 whereas in cotton and fibres from other plants DP values are usually in the range 800‐10 000 depending on the extraction method (Klemm et al., 2005). The formation of crystalline cellulose is dependent on interactions between the free hydroxyl groups of the molecule forming intra‐ and intermolecular bonds. In addition, the three dimensional structure of cellulose involves stacking of cellulose sheets through weak hydrogen bonds (C—H O) as well as hydrophobic 14 interactions (Jarvis, 2003). The different forms of cellulose define four types depending on the unit cell parameters of the crystals: cellulose I, II, III and IV. The different forms can be identified by their characteristic X‐ray diffraction patterns. Cellulose I occurs in two distinct allomorphs designated Iα and Iβ, which can both exist in a given sample. The ratio of the Iα and Iβ allomorphs depends on the origin of the cellulose: cellulose Iα is most abundant in bacteria and certain algae while the cellulose of plants mainly consists of the Iβ allomorph (Atalla and Vanderhart, 1984). Cellulose II can be formed irreversibly from cellulose I by two different processes: mercerization or regeneration. Mercerization includes swelling in aqueous sodium hydroxide and recrystallization of the cellulose. Regeneration involves dissolution of the cellulose, e.g. in N‐methylmorpholine‐N‐oxide, and subsequent precipitation and crystallization, as for example in the process of making film or rayon fibres (reviewed by Klemm et al., 2005). Cellulose II is more thermodynamically stable than cellulose I. In cellulose I, the chains have a parallel orientation where the reducing end of all chains point to the same direction as opposed to cellulose II, which is antiparallel (Klemm et al., 2005). Cellulose III can be prepared from cellulose I or II by different chemical treatments and cellulose IV is obtainable by heating cellulose III in glycerol (Pérez and Mazeau, 2005). Cellulose I, also called native cellulose, and cellulose IV are the only forms encountered in nature. Cellulose IV is considered as a disorganized form of cellulose I and occurs in plant primary walls and walls of some microorganisms, e.g. Oomycetes (Chanzy et al., 1979; Bulone et al., 1992; Helbert et al., 1997).

Figure 6. The cellulose chain is composed of D‐glucose residues linked through β‐(14) linkages, with a reducing end and a non‐ reducing end, referring to the different chemical reactivity of the chain ends. The repeating unit is sometimes considered to be cellobiose, because of the 2‐fold screw axis of the glucan chain and a 180° rotation between each glucose monomer.

4.2 Chitin Chitin is the second most abundant natural carbohydrate after cellulose. It is present in crustaceans, arthropods, fungi, protists and some Oomycetes. It can be found for instance in the cell walls of fungi, in amoeba cysts and in the exoskeleton of insects, shellfish and snails. As cellulose, chitin is biodegradable. Chitin has immunostimulatory effects, for instance it accelerates wound healing (Hirano, 1999). Therefore, one of the major application areas of chitin is the biomedical industry. In addition, chitin is important in many other sectors such as for instance in the food industry, and for the treatment of industrial pollutants (Hirano, 1999). The chemical reactivity of the amine groups of chitin makes it attractive for industrial applications after modification. The most important chitin derivative is chitosan, obtained after partial deacetylation of chitin under alkaline conditions. Sulfated chitin and chitosan have heparin‐like properties, are antibacterial and can chelate metal ions. They are promising as new biomaterials e.g. for drug delivery, as anti‐microbial agents and anticoagulants (Jayakumar et al., 2007).

Chitin structure Chitin is a linear homopolymer of β‐(14) linked N‐acetylglucosamine residues (Figure 7). Chitin, similarly to cellulose, contains a reducing and non‐reducing end. Chitin microfibrils 16 consist of aggregated chains, associated essentially by hydrogen bonds. There are two main types of crystalline chitin, the α‐ and β‐forms (Rinaudo, 2006). A third unusual crystal form of chitin exists, γ‐chitin, which can be found in some insects (Hudson and Smith, 1998). In α‐ chitin, the chitin chains are antiparallel, which means that the orientation of the reducing ends is alternating in the structure. The structure of β‐chitin is parallel, with all the reducing ends of the chitin chains pointing in the same direction. In γ‐chitin, the orientation of the chitin chains are antiparallel, with two chains in one orientation and one chain in the other orientation in alternating order (Hudson and Smith, 1998). The stability of α‐ and β‐chitin differs. α‐Chitin contains inter‐ and intrasheet hydrogen bonds whereas β‐chitin only contains intrasheet bonding, and hence β‐chitin has weaker interactions and is less stable than α‐chitin (Hudson and Smith, 1998). The α‐form is the most widespread form in nature and exists in arthropods, fungi and entamoeba. β‐chitin is less common and can be found in squid pens, some other marine animals (e.g. vestimentiferans), and the spines of certain diatoms (Kurita, 2006). In the native organism, chitin fibrils are covered with proteins. The structure of chitin varies depending on which organism it is extracted from. For instance, the degree of acetylation varies depending on the natural source of chitin and the extraction method. However, the degree of acetylation of extracted chitin from crab, shrimp and squid is around 90‐95 % (Kurita, 2006). Chitosan is obtained when the degree of chitin deacetylation reaches around 50 % (Rinaudo, 2006). Chitin can form a particularly strong material when associated to other compounds, such as proteins and minerals, in the form of a composite (e.g. crab shells). Pure chitin also has remarkable mechanical properties comparable to cellulose, which is useful for instance for the preparation of surgical threads (Dutta et al., 2004).

Figure 7. Structure of a fully acetylated chitin chain. Similar to cellulose, the chitin chain contains a reducing and non‐reducing end (see text for explanation).

5. CELLULOSE AND CHITIN BIOSYNTHESIS IN DIFFERENT ORGANISMS

5.1 Catalytic subunits of cellulose and chitin synthases Cellulose synthases (CesAs) and chitin synthases (Chs) are responsible for the biosynthesis of cellulose and chitin in nature. In this section, their general characteristics are presented. In addition to describing the aspects related to the glycosyltransferase activities of these enzymes, a section briefly covers the additional domains that are found in some of these proteins. Pleckstrin homology (PH) domains are of special interest, since they are part of the present investigation; they are described more thoroughly.

5.1.1 Classification and properties of cellulose and chitin synthases Glycosyltransferases are classified into distinct sequence‐based families (Coutinho et al., 2003) described in the carbohydrate‐active enzyme (CAZy) database (www.cazy.org; Coutinho and Henrissat, 1999), which currently comprises 92 families of glycosyltransferases (GTs). CesAs and Chs belong to GT family 2 (GT2), one of the largest of the GT families in CAZy. GT2s use an inverting mechanism (see 5.1.2, Inverting mechanism of GT2s) and include both processive and non‐processive enzymes. Processive GTs transfer multiple sugar residues to the growing polymer whereas non‐processive GTs transfer a single sugar residue to the acceptor (Saxena et al., 1995). CesA and Chs are both processive GTs, and are related

18 to other processive GT2s such as hyaluronan synthase by sequence similarity and probably partially shared structures (Charnock et al., 2001).

Conserved motifs of CesAs and Chs Processive GTs belonging to family 2 contain two conserved domains, A and B (Figure 8). Most non‐processive GTs from the same family contain only domain A, suggesting that the conserved aspartic acid and the QXXRW (Q(R/G)RRW in Chs enzymes) motif of domain B are involved in the processivity of the enzyme. However, some enzymes that possess domain B and the QxxRW motif are known to be non processive (www.cazy.org). Mutagenesis of the two conserved aspartic acid residues in domain A and that in domain B, as well as mutagenesis of the Q, R and W residues in the QXXRW motif in the CesA from the bacterium Gluconacetobacter xylinus (formerly Acetobacter xylinum), has shown that these amino acids are necessary for enzyme activity (Saxena and Brown, 1997; Saxena et al., 2001). All known CesA and Chs genes encode transmembrane proteins. Plant CesAs are usually predicted to have six or more transmembrane helices, two located at the N‐terminal end and the remaining ones at the C‐terminus. In the Chs proteins, all transmembrane helices are located in the C‐terminus and their N‐terminal domain is completely soluble. Eukaryotic CesAs have additional domains at their N‐terminus. Plant CesAs have longer N‐terminal ends than bacterial CesAs (Figure 8). Their N‐terminal ends include a conserved zinc finger domain, which contains four repeated CXXC motifs. This domain has been shown to be involved in the homo and hetero dimerization of cotton CesAs in vitro and in a yeast two‐hybrid system (Kurek et al., 2002), but the function in vivo is not known. Animal CesAs contain a glycosidehydrolase family 6 cellulase domain (Nakashima et al., 2004). However, the catalytic residues are mutated and the cellulase domain is probably not hydrolytic but fulfils some other unknown function. PH domain is another example of an N‐terminal CesA domain and exists only in the Oomycete CesA isoforms 1, 2 and 4 (figure 8; Grenville‐Briggs et al., 2008; Fugelstad et al., 2009). The function of this domain has been studied as a part of the present investigation. Specific N‐terminal domains can be found in Chs proteins as well. One example is the Chs class V and VII of some fungi, which contain a domain similar to the myosin motor‐like domain (Takeshita et al., 2005). These Chs localize near actin rich structures at the hyphal tips and septum in vivo, presumably via the myosin motor‐like

19 domain. In this thesis work is presented in paper III which demonstrates the presence of a microtubule‐interacting and trafficking domain at the N‐terminus of Chs1 and Chs2 from S. monoica.

Figure 8. Domain arrangement of CesAs from plants, Oomycetes and bacteria. Highlighted in red box: domain A of processive and non‐processive GT2s. Highlighted in green box: domain B of processive GT2s. PH: Pleckstrin homology domain. See text for further explanation.

5.1.2 Inverting mechanism of GT2s There is no crystal structure available for any processive glycosyltransferase from the GT2 family and the reaction mechanism of CesA is poorly understood. The inverting mechanism is a one‐step reaction where the configuration of the anomeric carbon is inverted with respect to the sugar donor. CesAs, Chs and other inverting GTs form β‐linkages and use an α‐ linked donor substrate, such as for instance uridine‐diphosphate‐α‐D‐glucose (UDP‐glucose) in the case of CesA and uridine‐di‐phospho‐α‐N‐acetyl‐D‐glucosamine (UDP‐GlcNAc) for chitin synthase. The reaction involves a nucleophilic substitution at the anomeric carbon of the nucleotide‐sugar (Figure 9). According to the scheme, a catalytic residue, most likely a

20 carboxylate, is acting as a base by deprotonating a hydroxyl group from the non‐reducing end of the acceptor, thereby activating it (Saxena et al., 1995; Charnock et al., 2001). A second carboxylate is thought to be involved in the coordination of a metal ion, e.g. Mg2+, and the nucleotide‐sugar (Figure 9). The activated acceptor thereafter attacks the anomeric carbon of the coordinated nucleotide‐sugar, forming a β‐glycoside bond and releasing the nucleotide moiety of the donor. There are only two published three‐dimensional structures determined from the GT2 family, the structure of the bacterial nucleotide‐sugar transferase SpsA from Bacillus subtilis (Charnock and Davies, 1999) and the structure of teichoic acid polymerase TagF from Streptomyces epidermis (Strynadka et al., 2010). The function and sugar donor of SpsA are not known but the structure has been used to discuss the possible organization of the catalytic site of CesA (Charnock et al., 2001). The TagF is a non‐processive glycosyltransferase which catalyses the addition of a glycerolphoshate moiety to a lipid disaccharide acceptor (Brown et al., 2009). The other mechanism for glycosyl transfer is the retaining mechanism, where the configuration of the anomeric carbon of the sugar donor is retained. This involves the formation of a glycosyl‐enzyme intermediate and the subsequent addition of the acceptor (Sinnott, 1990; Davies and Henrissat, 1995; Saxena et al., 1995; Lairson et al., 2008). A typical example of a GT using this mechanism is glycogen synthase, which forms an α bond from a UDP‐glucose molecule in which the glucose is in the α configuration. Whether the chain elongation of cellulose takes place at the reducing or non‐ reducing end is debated. Electron microdiffraction‐tilting of stained reduced ends of G. xylinus cellulose, showed that the reducing ends of the growing cellulose chain point away from the bacterium (Koyama et al., 1997). This provides some evidence that the cellulose molecule is elongated from the non‐reducing ends of the growing chains in G. xylinus. Other experimental data, based on 14C‐pulse and chase experiments with radioactive D‐glucose and UDP‐glucose support an elongation from the reducing end (Han and Robyt, 1998). Transmission electron microscopy experiment on β‐chitin, still attached to the catalytic center of the enzyme and labeled at its reducing end, have shown that the chitin chain is synthesized with its reducing end protruding away from the catalytic center (Imai et al., 2003) supporting elongation at the non‐reducing end of chitin. In the case of other GT2s, like the hyaluronan synthase, there is strong experimental evidence that the elongation of the chain occurs from the non‐reducing end (DeAngelis, 1999). To summarize, most results from

21 studies with CesA and related glycosyltransferases support polymerization of cellulose and other linear homopolysaccharides at the non‐reducing end.

Figure 9. Proposed mechanism of inverting GTs: a nucleophilic substitution at the anomeric carbon of the nucleotide‐sugar leads to the formation of a β‐linkage from an α‐linked donor. A carboxylate is thought to coordinate a metal ion associated with the nucleotide‐sugar (reprinted with permission from Charnock et al., 2001).

5.1.3 Pleckstrin homology domains In the present work, a functional study of the PH domain from S. monoica CesA2 is presented. In this section, the general characteristics and functions of PH domains are presented.

What are the PH domains? The name pleckstrin homology domain was adopted in 1993 (Mayer et al., 1993) after the human protein pleckstrin, a major substrate of protein kinase C in platelets (Tyers et al., 1988). The pleckstrin protein contains two PH domains (Haslam et al., 1993). PH domains are around 100 amino acids long and exist mainly in eukaryotes. The first structure of a PH domain was solved in 1994 by NMR (Yoon et al., 1994). Many other PH domain structures have been determined after that, such as those of β‐spectrin, dynamin 1, son of sevenless1

(Sos1), G protein‐coupled receptor kinase 2 (GRK2), phospholipase Cδ1 (PLC δ1), Brutons

22 tyrosine kinase (Btk) and insulin receptor susbtrate 1 (IRS1) (Downing et al., 1994; Macias et al., 1994; Ferguson et al., 1995; Hyvonen and Saraste, 1997; Koshiba et al., 1997; Zheng et al., 1997; Fushman et al., 1998; Dhe‐Paganon et al., 1999). Rather than the amino acid sequence, it is the tertiary structure of the domain that is conserved (Lemmon and Ferguson, 2000). The general PH domain fold consists of a seven β‐sheet core with a C‐terminal α‐helix (Figure 10).

Figure 10. Structure of the C‐terminal PH domain of the tandem PH domain‐containing protein‐1 (TAPP1), a human protein. TAPP1 interacts with a citrate molecule (pdb id: 1EAZ). Structure made in PyMOL.

Functional characteristics of PH domains The function of most known PH domains is still unclear. Basically, all investigated PH domains bind to phosphoinositides, although most of them in a non‐specific manner and with low affinity (Lemmon, 2004). Phosphatidylinositides (PtdIns, Figure 11) are important in cell signaling and regulation. They occur in minor amount in the cell membrane and their membrane and intracellular levels can change fast. The PtdIns act as downstream signaling molecules, in some cases through interactions with PH domains. PH domains can often be

23 found in signaling molecules, for instance in guanine exchange factors, cytoskeletal proteins (such as spectrin), putative singaling adaptor proteins (such as IRS‐1) and in proteins involved in cellular membrane transport (such as dynamin) (Allam and Marshall, 2005). The polar surface of the PH domain structure exhibits a net positive charge and is believed to cause the sometimes non‐specific binding to PtdIns, which are negatively charged. However, about 10 % of the PH domains in human proteins show very specific and high affinity binding to certain PtdIns. These PH domains usually share a phosphoinositide‐binding motif, located in the two first β‐sheets, containing amino acids with basic properties. Some of these basic residues have been shown to interact directly with the phosphates in the phosphorylated PtdIns (Isakoff et al., 1998; Kavran et al., 1998; Lemmon and Ferguson, 2000; Lemmon, 2008). Most studied PH domains are from mammal proteins, but there are PH domains in other organisms as well. For instance, in the yeast S. cerevisiae there are around 30 PH domain containing proteins in the genome (Lemmon, 2008). Phosphoinositide kinase‐3 (PI3‐kinase) belongs to a group of phosphoinositide 3’ kinase related‐kinases (PIKK). PIKKs have a role in PtdIns metabolism and are involved for instance in stress responses. There are PIKKs in Oomycetes, but their function are not well characterized (Meijer and Govers, 2006; Judelson and Ah‐Fong, 2010). In human cells, PI3‐ kinase is responsible for the generation of phosphatidylinositol(3)‐phosphate (PtdIns(3)‐P), phosphatidylinositol(3,4)di‐phosphate (PtdIns‐(3,4)‐P2) and phosphatidylinositol(3,4,5)tri‐ phosphate (PtdIns‐(3,4,5)‐P3) upon stimulation (Leevers et al., 1999). These PtdIns can also be generated in PI 3‐kinase independent pathways. Phosphatidylinositol(3,5)di‐phosphate

(PtdIns‐(3,5)‐P2) is obtained by phosphorylation of PtdIns(3)‐P, for instance by a phosphoinositide‐interacting 5‐kinase (PI 5‐kinase). An example of a PH domain which binds to a PI‐5‐kinase product is the PH domain of PLC δ1. This PH domain was one of the first to be functionally characterized (Lemmon et al., 1995) and it binds preferably to PtdIns‐(4,5)‐P2

(Ferguson et al., 1995). For some proteins containing PH domains, like PLC δ1, the PH domain is crucial for membrane targeting of the protein (Paterson et al., 1995). Many PH domains that bind specifically to PtdIns bind to PI3‐kinase products, like the PH domain of Btk, which binds specifically to PtdIns‐(3,4,5)‐P3 (Baraldi et al., 1999). The C‐terminal PH domain of

DAPP1 is another example, and binds both PtdIns‐(3,4,5)‐P3 and PtdIns‐(3,4)‐P2 (Dowler et al., 1999). Other PH domains are more promiscuous and bind to all phosphorylated PtdIns in an unspecific way and with less affinity, like the ones of the oxysterol‐binding protein and β‐

24 spectrin. They bind PtdIns‐(3,4,5)‐P3 and PtdIns‐(4,5)‐P2 with low affinity (Rameh et al., 1997). Many PH domains that interact with PtdIns are largely located in the cytosol until stimulation of cells. Upon stimulation, a phosphoinositide kinase activity increases PtdIns levels in the membrane, leading to the targeting of the protein containing the PH domain to the membrane (Isakoff et al., 1998). Two examples of such signal dependent membrane localized proteins are the tandem PH domain protein 1 (TAPP1, Figure 10) and the general receptor for phosphoinositides‐1 (Grp1). The C‐terminal PH domain of TAPP1 remains mainly cytosolic in human B‐lymphoma cells until stimulation of the B‐cell antigen receptor (BCR) (Marshall et al., 2002). When the BCR is stimulated, the C‐terminal PH domain of TAPP1 is targeted to the plasma membrane, where it locates in filamentous actin (F‐actin) rich ruffles. Grp1 is located in the cytosol in human embryonic kidney cells. Upon insulin stimulation,

Grp1 is targeted to the membrane, due to the increased levels of PtdIns(3,4,5)P3 (Gray et al., 1999). Other observed binding partners of some PH domains are actin, protein kinase C and βγ subunits of hetrotrimeric G proteins (Touhara et al., 1994; Yao et al., 1994; Yao et al., 1999). However, the binding partners of most of the PH domains are still unknown.

Figure 11. Structure of PtdIns. R: fatty acid chain.

5.2 Cellulose biosynthesis

5.2.1 Terminal complexes responsible for cellulose biosynthesis Cellulose synthesis takes place at the plasma membrane in so‐called terminal complexes. In bacteria, tunicates and most algae, the terminal complexes are organized as linear rows of catalytic subunits, single or multiple, depending on the species considered (figure 12 A; Brown, 1996). In plants and some algae, the terminal complexes are organized in hexagonal structures called rosettes (Figure 12 B). Freeze‐fracture experiments combined with immunogold labelling, demonstrated for the first time the occurrence of the cellulose synthase catalytic subunit (CesA) in the rosettes (Kimura et al., 1999).

Bacteria Algae Plants/Algae

A B

Figure 12. Schematic representation of terminal complexes. A) The terminal complexes are linearly organized in single rows in bacteria and multiple rows in most algae. In plants (and some algae) the terminal complexes are organized as hexagonal or octagonal (not shown) structures designated as rosettes (after Brown, 1996). B) Freeze‐fracture of the plasma membrane from a tracheary element of Zinnia elegans. Main micrograph: magnification 222 000 times. Inset magnification: 504 545 times (reprinted with permission from Delmer, 1999).

5.2.2 Cellulose biosynthesis in bacteria Bacterial cell walls are essentially composed of peptidoglycan and lack cellulose. Thus, bacterial cellulose exerts other functions than building the cell wall. In bacterial species that produce cellulose, the polymer is always secreted as an extracellular polysaccharide either in the form of ribbons composed of packed microfibrils (e.g. G. xylinus and Agrobacterium tumefaciens; Ross et al., 1991), or inserted into slime tubes, sheats or extracellular slimy materials (e.g. cyanobacteria; Nobles et al., 2001) or biofilms (Pseudomonas species and Escherichia coli; Ude et al., 2006; Matthysse et al., 2008). It is believed that the cellulose ribbons produced by G. xylinus are used as a raft by this aerobic bacterium to float at the water‐air interface. A. tumefaciens uses cellulose to attach to the host (Matthysse et al., 1981) and this has also been reported for E. coli (Matthysse et al., 2008). Bacteria have been the preferred model system to study cellulose biosynthesis during the 90’s and the most studied bacterium has been G. xylinus (reviewed by Römling, 2002). In G. xylinus, cellulose is synthesized by linear terminal complexes and the cellulose chains are secreted outside the cells, forming bundles that assemble into a loose ribbon of fibrils containing approximately 1000 glucan chains. UDP‐glucose is the substrate for cellulose synthesis in G. xylinus (Glaser, 1958). Cyclic diguanylic acid (c‐di‐GMP) and Mg2+ ions regulate the rate of synthesis, while lipids and protein‐linked cellodextrins have been proposed to act as intermediates (Swissa et al., 1980). The catalytic subunit of the bacterial CesA was identified by photolabelling with the photoaffinity probe 5‐azido‐UDP‐glucose, which labelled a UDP‐glucose‐binding polypeptide of 83 kDa (Lin et al., 1990). The genes involved in cellulose biosynthesis were first identified in G. xylinus by complementation of a cellulose deficient mutant (Wong et al., 1990). Cellulose biosynthesis is controlled by an operon containing four genes including that encoding the CesA catalytic subunit. The operon is designated as bcs or acs depending on the authors (hereafter the name bcs is used), and accordingly the CesA catalytic subunit corresponds to the bcsA or acsA gene (Saxena et al., 1990; Wong et al., 1990). The bcsB gene encodes a polypeptide probably involved in binding c‐di‐GMP, and thus functions as a regulator of cellulose synthesis (Mayer et al., 1991). Mutational studies propose that bcsD is involved in the crystallization of cellulose during cellulose biosynthesis (Saxena et al., 1994). Cellulose content in bscD mutants decreased to only around 9 % compared to the wild‐type control in vivo (Hu et al., 2010). The 3D structure of the bcsD protein was recently obtained together with cellopentaose (Hu et al., 2008; Hu

27 et al., 2010) revealing an octamer structure with four inner passageways, presumably for extrusion of the cellulose chain across the outer cell membrane. The function of bcsC remains unknown. A cellulase from the CAZy glycoside hydrolase family 8, located upstream of the bcs operon, is necessary for cellulose biosynthesis in G. xylinus (Standal et al., 1994). A homologue of the G. xylinus cellulase has been found in the A. tumefaciens celA operon (Matthysse et al., 1995a). There is limited information about cellulose biosynthesis in other bacteria than G. xylinus and A. tumefaciens.

5.2.3 Cellulose biosynthesis in higher plants It was not until 1996 that the first plant CesA gene was isolated and sequenced in cotton (Gossypium hirsutum). The sequence was identified based on sequence similarity with the bacterial bcsA of sequenced cDNA clones from developing cotton fibers, in which cellulose synthesis is high during the formation of secondary cell walls (Pear et al., 1996). It was demonstrated that a recombinantly expressed fragment of GhCesA1, the first CesA homologue of bcsA identified in plants, binds UDP‐glucose, which is also the substrate of plant CesAs. Radial swelling mutants (rsw) in Arabidopsis thaliana (hereafter Arabidopsis) exhibit a reduced cellulose content, accumulation of noncrystalline (14)‐β‐glucan, disassembly of the CesA rosettes, and widespread morphological abnormalities (Baskin et al., 1992; Arioli et al., 1998). The involvement of CesA in cellulose biosynthesis in plants was confirmed by the complementation of a rsw Arabidopsis mutant with the wild‐type allele, a homologue of GhCesA (Arioli et al., 1998). To date, the only evidence of the catalytic activity of CesA proteins is indirect and based exclusively on in planta studies.

Priming of cellulose polymerization Cellulose synthesis in A. tumefaciens requires a glycolipid intermediate (Matthysse et al., 1995b), and some observations support the involvement of lipid‐linked intermediates also in cellulose synthesis in G. xylinus (Han and Robyt, 1998). In plants, a sitosterol‐β‐glucoside has been suggested to function as a primer for cellulose synthesis (Peng et al., 2002). However, mutants deficient in the enzymes that encode UDP‐glucoside:sterol glycosyltransferases are not impaired in cellulose biosynthesis, but show a decrease in cuticle polymers (DeBolt et al.,

2009). Sterol‐glucosides in these mutants are almost completely depleted, although residual sterol‐glucosides could still be detected and it cannot be ruled out that these might be enough to function as primers for cellulose biosynthesis. Arabidopsis mutants completely deficient in sterol biosynthesis are however impaired in cellulose biosynthesis while they have normal pectin and total sugar content (Schrick, 2004). These results indicate indeed a role for sterols in cellulose biosynthesis, but the function of a possible primer needs to be further investigated (Figure 13, Cellulose synthesis regulation and proteins involved in cellulose biosynthesis).

Formation of the cellulose synthase complex in plants Plant CesAs have numerous homologues in all plant species: 10 have been identified in the small annual plant Arabidopsis (Henrissat et al., 2001) while 18 occur in the perennial tree Populus trichocarpa (Djerbi et al., 2005). It has been shown that some CesAs are associated to primary cell wall biosynthesis while others seem to be linked to secondary cell wall formation. These conclusions arise from observations in Arabidopsis mutants (reviewed in Endler and Persson, 2011) and expression profiling of CesA genes in other plants (e.g. Djerbi et al., 2004; Suzuki et al., 2006). Three unique primary cell wall associated Arabidopsis CesAs, namely AtCesA1, AtCesA3 and AtCesA6, are needed to form a functional complex. AtCesA5 and AtCesA2 are partially redundant with AtCesA6 (Persson et al., 2007). In addition, AtCesA3 and AtCesA6 co‐immunoprecipitate in vitro (Desprez et al., 2007). Bimolecular fluorescence complementation (BiFC), using chimeric CesAs with partial yellow fluorescent proteins (YFP), is a way to study protein‐protein (CesA‐CesA) interactions in vivo. With this technique, it was shown that AtCesA1, AtCesA3 and AtCesA6 can form homo‐ and heterodimers with each other in a Nicotiana benthamiana leaf system (Desprez et al., 2007). AtCesA8, AtCesA7 and AtCesA4 seem specific to secondary cell walls. They co‐ immunoprecipitate in vitro and AtCesA4 is needed for the formation of the complex (Taylor et al., 2000; Taylor et al., 2003). The secondary cell wall associated CesAs from Arabidopsis are able to form heterodimers with each other in a split ubiquitin yeast two‐hybrid system (Timmers et al., 2009). This was confirmed by BiFC (Timmers et al., 2009). However, only AtCesA4 can form homodimers. In poplar differentiating xylem, two separate types of cellulose synthase complexes have been identified by co‐immunoprecipitation. One type

29 contains six CesAs corresponding to Arabidopsis primary CesAs while the other type comprises five CesAs corresponding to Arabidopsis secondary CesAs, respectively (Song et al., 2010). As mentioned previously, the terminal complex rosette in plants consists of six globules (Figure 12). Each globule is typically described as containing six CesA catalytic subunits, so that the whole CesA complex produces cellulose elementary microfibrils composed of 36 chains (Figure 13; Delmer, 1999). However, the actual number of catalytic subunits in a rosette has never been experimentally determined and the definite composition of the rosette is not clear. The stoichiometry of CesAs in the complex is of great importance to obtain normal cellulose biosynthesis in Arabidopsis. However, redundancies exist between some CesA isoforms (Persson et al., 2007; Endler and Persson, 2011). The zinc finger domain of plant CesAs has been proposed to be responsible for the interaction between CesAs, since they can form homo and heterodimers in vitro (Kurek et al., 2002), but this needs to be confirmed in vivo. It can be concluded that certain CesAs are more important for secondary cell wall synthesis and others for primary cell wall synthesis. It should however be noted that the majority of the studies have been made in Arabidopsis, which is an annual plant that contains very little vascular tissue. Hence, it is difficult to draw general conclusions about secondary and primary cell wall associated CesAs from this plant model.

Figure 13. Hypothetical model for cellulose biosynthesis in plants (after Guerriero et al., 2010). The model shows a CesA catalytic subunit in the plasma membrane. Several key aspects related to the cellulose biosynthesis process are poorly understood and indicated with red question marks in the figure: the role of the endoglucanase Korrigan and its physical association to the complex, the involvement of a primer, the mode of translocation of the nascent cellulose chain across the plasma membrane, the role of sucrose synthase (SuSy), the role of the cytoskeleton and the stoichiometry of the complex.

Regulation of cellulose biosynthesis and proteins involved in cellulose formation Korrigan, KOBITO1 and COBRA are examples of genes involved in cellulose biosynthesis in Arabidopsis (Nicol et al., 1998; Schindelman et al., 2001; Pagant et al., 2002). A protein similar to Korrigan also occurs in poplar and is up‐regulated during secondary cell wall formation (Rudsander et al., 2003). Korrigan is a glycoside hydrolase family 9 membrane‐ bound β‐1,4‐endoglucanase involved in cellulose biosynthesis in Arabidopsis. However, not much is known about the precise function of any of the Korrigan gene products and so far none of them have been found to associate directly with the cellulose synthase complex (Figure 13). The involvement of sucrose synthase in cellulose biosynthesis has been proposed. Recently, the association of a sucrose synthase‐like protein to the membrane‐ bound cellulose synthase complex from bean was reported (Fujii et al., 2010). Contradictory with these results, an Arabidopsis mutant devoid of any functioning sucrose synthase in the whole plant except from phloem tissues, contains normal levels of cellulose (Barratt et al., 2009). These results question the role of SuSy in cellulose biosynthesis (Figure 13). The role of the cytoskeleton in cellulose biosynthesis has long been debated (Lloyd and Chan, 2008). Lately, the use of YFP‐CesA6 has allowed the visualisation of the CesA complex associated to microtubule bundles in Arabidopsis (Paredez et al., 2006). However, the movement of the CesA complex in the plasma membrane is no longer thought to be driven by microtubule dynamics, but it is rather the polymerisation of cellulose itself that is believed to propel the cellulose synthase complex in the plasma membrane (Endler and Persson, 2011). Actually, the cellulose synthesis activity alone has been shown to influence microtubule organization (Paredez et al., 2008). In the experiment, microtubule organisation was shown to be disturbed in Arabidopsis mutants with impaired cellulose synthesis activity, or in wild type cells treated with the cellulose synthase inhibitor isoxaben. A model where microtubule bundles initiate the track of the cellulose synthase complex has been proposed (Figure 14; Mutwil et al., 2008; Crowell et al., 2010). This model is based on results that have supported a role for cortical microtubules in the initial guiding of the cellulose synthase complex in the plasma membrane, as visualized by using GFP‐labelled CesA3 and a mCherry‐ labelled microtubule‐associated protein or tubulin in Arabidopsis (Crowell et al., 2009; Gutierrez et al., 2009). In addition, there have been several observations of structures believed to be Golgi bodies delivering cellulose synthase complexes to the membrane, by docking to microtubules (Crowell et al., 2009; Gutierrez et al., 2009). The actin cytoskeleton

32 is assumed to be responsible for the global distribution in the cell of the Golgi bodies carrying the cellulose synthase complexes (Crowell et al., 2009; Gutierrez et al., 2009). To conclude, most studies point towards a relationship between microtubules and the cellulose synthase machinery and a global role for the cytoskeleton in transport and delivery of the cellulose synthase complex to the plasma membrane. A microtubule‐associated protein, MAP20, is upregulated during poplar secondary cell wall synthesis (Rajangam et al., 2008). MAP20 was shown to bind microtubules and the cellulose synthase inhibitor DCB in vitro, and was co‐localised with microtubules when expressed as a YFP‐fusion protein in N. benthamiana leaves. MAP20 might serve as a link between the cellulose synthase complex and the microtubules in secondary cell walls. However, in Arabidopsis it is still unclear if the cytoskeleton is linked to the CesA complex directly, or if there are one or several intermediate proteins. If it is the case, the mode of regulation of this interaction is unknown.

Figure 14. Hypothetical model describing the possible role of microtubules in cellulose biosynthesis (adapted and reprinted with permission from Mutwil et al., 2008). The CesA complex (in red) is transported from the Golgi stacks via the trans‐Golgi network (TGN) to the plasma membrane, where the microtubules (MT) sometimes initiate a track for the CesA complex movement. Inset: average of YFP‐tagged CesA6 (green) and CFP‐tagged tubulin (red) in an Arabidopsis etiolated hypocotyl cell (reprinted with permission from Paredez et al., 2006).

Cellulose synthase is part of the dynamic cell wall synthesis machinery, a system that is very tightly regulated, since building of the cell wall is part of the differentiation process of plant cells. For instance, during secondary cell wall synthesis a thick and rigid cell wall is deposited after cells have elongated. When the secondary cell wall is deposited cells undergo apoptosis, meaning that a high cellulose synthesis must happen at a precise time not to rigidify cells before they have expanded. The regulation is also reflected in attempts to synthesize cellulose in vitro, which mostly leads to callose synthesis, indicating that the cellulose synthase machinery is inactivated rapidly by some unknown mechanism. The half life of cellulose synthase in cotton fibers is only around 30 minutes, indicating a rapid turnover (Jacob‐Wilk et al., 2006). However, very little is known about the regulation of the

CesA complex. Phosphorylation seems to be part of the regulation process. A phosphoproteomic study in Arabidopsis revealed several phosphorylation sites in several CesAs, mostly located in the N‐terminus (Nühse et al., 2004). Arabidopsis CesA1 has several phosphorylation sites in the N‐ and C‐terminal plant‐specific regions. All AtCesA1 phosphorylation sites have been point mutated to alanine or glutamate to eliminate or mimic phosphorylation respectively (Chen et al., 2010). The constructs were used to complement an Arabidopsis mutant for AtCesA1. The results indicate that phosphorylation is involved in the regulation of AtCesA1 in rapidly elongating cells, since effects linked to the different substitutions could be seen on growth (Chen et al., 2010). Certain kinases have been shown to influence cellulose synthesis in Arabidopsis. For instance, two leucine‐rich repeat receptor kinases, FEI1 and FEI2, are important for the synthesis of crystalline cellulose during anisotropic cell expansion (Xu et al., 2008). A receptor‐like kinase, THESEUS1, expressed in expanding cells and vascular tissue, partially restores the phenotype of CesA6 mutants in Arabidopsis (Hematy et al., 2007). When attempting to overexpress the poplar secondary cell wall related CesA8, silencing of the corresponding native gene as well as the transgene was obtained, leading to a massive decrease in cellulose content in the transgenic wood (Joshi et al., 2011). This might indicate some sort of disturbed regulation of the stochiometry upon introducing the transgene. In summary, phosphorylation as well as the stoichiometry of the complexes are probably involved directly in the regulation of the cellulose synthase machinery. The observations from the phosphorylation mutations in AtCesA1 are interesting and should be confirmed in another system. It is also needed to investigate whether other CesA homologues contain phosphorylation sites and how these regulate the cellulose synthase complex. In addition, the targets of the studied kinases involved in cellulose synthesis are unknown. It could be CesA proteins themselves, other proteins involved in the cellulose synthase machinery or any other regulating protein. The cellulose synthase complex is sensitive to perturbations, as indicated from the overexpression of a poplar CesA that lead to decreased cellulose content (Joshi et al., 2011) and from the loss of activity upon extraction of the complex in in vitro experiment (Delmer, 1999; Jacob‐Wilk et al., 2006).

Cellulose synthase‐like proteins In addition to CesAs, plants contain so‐called cellulose synthase‐like (Csl) proteins (Figure 15). These proteins belong to the glycosyltransferase family 2 and share the conserved D, D, D, QXXRW motifs with CesAs. However, they do not contain the zinc finger motif of CesAs (see section 5.1.1; Conserved motifs of CesAs and Chs). At least some of the Csl proteins are predicted to be involved in the synthesis of non‐cellulosic cell wall carbohydrates and are believed to be active in the Golgi apparatus. However, only a few Csl proteins have been functionally characterized. CslF and CslH are involved in the synthesis of the monocot‐ specific β‐(13, 14)‐glucan (Burton et al., 2006; Doblin et al., 2009). Experimental data indicate that proteins of the CslA family synthesize β‐mannans and β‐glucomannans (Dhugga et al., 2004; Liepman et al., 2005; Burton et al., 2006; Goubet et al., 2009). A heterologously expressed CslC has been shown to synthesize β‐(14)‐glucans in vitro and is probably involved in the formation of the xyloglucan backbone (Cocuron et al., 2007). The disruption of a CslD in Arabidopsis leads to a decreased growth of the roots and stems, and lower levels of xylan in stems (Bernal et al., 2007). When using microsomes in in vitro synthesis of xylan and homogalacturonan, the activity was decreased in the mutant compared to wild type. These results indicate a role in xylan biosynthesis for the CslD gene in Arabidopsis. CslB remains unchararacterised as well as most members of the different Csl families except those mentioned above.

Figure 15. The CesA/Csl superfamily. Figure adapted from Carpita (2011) (www.plantphysiol.org; Copyright American Society of Plant Biologists).

5.2.4 Cellulose biosynthesis in Oomycetes As summarized in the previous section, there are still multiple questions to solve to fully understand the molecular mechanisms of cellulose biosynthesis in higher plants. However, even less is known about the formation of cellulose in Oomycetes. Sequence information from Oomycetes has not been available until recently. Therefore, most genes encoding the catalytic subunits of carbohydrate synthases in these microorganisms are not characterized. No CesA terminal complex has been observed in Oomycetes, as opposed to plants, bacteria and algae. In this thesis, we report the identification of a family of CesA genes based on the

37 work on two different Oomycete species. After our studies were published, a few other investigations on cellulose synthesis in Oomycetes have been conducted, showing the interaction of the cellulose synthase putative catalytic subunit with the drug mandipropamid (Blum et al., 2010a; Blum et al., 2010b). However, to date the Oomycete cell wall composition and in vitro characterization of cellulose synthase activity have been studied more than the molecular biological aspects of cellulose synthesis. The results from in vitro synthesis studies are summarized in sections 5.2.6 (Biochemical approaches to study cellulose synthesis in vitro and 5.4 (Inhibitors of glycosyltransferase activities). The results from the work included in this thesis are summarized in chapter 6 (Present investigation) and presented in detail in papers I, II and III.

5.2.5 Cellulose biosynthesis in other organisms Cellulose synthesis occurs in organisms other than those discussed in this thesis. However, the knowledge on cellulose formation in these organisms is rather scarce. Some CesA genes have been identified, and sometimes their function has been characterized. Examples are the CesAs from the algae Valonia and Microdictyon (Brown, 1985), slime molds of the genus Dictyostelium (Blanton et al., 2000) and amoebas of the genus Acanthamoeba (Anderson et al., 2005). Cellulose synthesis also occurs in a group of marine animals, the tunicates, and the corresponding CesA genes have been identified (Matthysse et al., 2004; Nakashima et al., 2004). In tunicates, cellulose is part of the exoskeleton of the animals.

5.2.6 Biochemical approaches to study cellulose synthesis in vitro Cellulose synthesis has been most studied in vitro in the bacteria G. xylinus and A. tumefaciens (Ross et al., 1991; Römling, 2002). In vitro synthesis of cellulose from UDP‐ glucose using cell free extracts of G. xylinus as a source of enzyme was reported for the first time in the 1950’s (Glaser, 1958). The cellulose synthase from G. xylinus is rather well characterized, but has never been expressed recombinantly and purified in an active form. The genes encoding plant and other CesAs have been identified 10 to 20 years ago, but the characterization of the corresponding activities in vitro has been difficult, essentially because of the high instability of the CesA complex in cell free extracts. In addition, in vitro synthesis

38 of cellulose using plant enzymes most often leads to very low yields of cellulose while β‐ (13)‐glucan is a major in vitro product, thereby complicating cellulose characterization. Plant β‐(13)‐glucan, also called callose, is normally produced in small amount by plant cells during cell division and the formation of phloem sieve plates. It is synthesized in higher amounts in specialized cells, e.g. in pollen mother cells and pollen tubes (Stone and Clarke, 1992). The production of callose is also triggered by different types of stress, for instance wounding or infection (Stone and Clarke, 1992). It has been suggested that the same enzyme could be responsible for the synthesis of both callose and cellulose in plants (Delmer, 1999). However, homologues of the yeast FKS protein, a putative β‐(13)‐glucan synthase catalytic subunit (Douglas et al., 1994), have been found in pollen tubes of Nicotiana alata (Brownfield et al., 2007), in barley (Hordeum vulgare) (Li et al., 2003), Arabidopsis (Hong et al., 2001) and cotton (Cui et al., 2001). These proteins belong to GT family 48 and do not exhibit the D, D, D, QXXRW motifs conserved in CesAs. In addition, GT2s resembling CesAs have been shown to synthesize β‐(13)‐glucan in Agrobacterium (Stasinopoulos et al., 1999). The callose produced normally during cytokinesis could be synthesized by a different enzyme than that responsible for the formation of the callose produced upon wounding or during in vitro extraction of membrane proteins (Saxena and Brown, 2000). Many attempts to improve the yields of in vitro synthesis of cellulose have been made, including separation techniques such as sedimentation in glycerol gradients of the protein extracts and the use of different detergents (Colombani et al., 2004). Some improvements have been achieved with larger proportions of cellulose produced in the case of the cotton and blackberry cellulose synthesis systems (Kudlicka et al., 1996; Lai‐Kee‐Him et al., 2002). Membrane preparations and detergent extracts of membranes from S. monoica have been used as a source of enzyme for in vitro synthesis of β‐(14) and β‐(13)‐glucans in the presence of UDP‐glucose. The products synthesized in vitro have typically been identified using specific cellulases and β‐(13)‐glucanases (Fèvre and Rougier, 1981; Bulone et al., 1990). The data showed that cellulose and β‐(13)‐glucan are co‐synthesized, but the proportion of cellulose is higher when the assay is performed in the presence of lower substrate concentrations (10 μM UDP‐glucose instead of 1mM) and 8 mM MgCl2. Microfibrillar glucans have been observed by transmission electron microscopy after in vitro

39 synthesis together with associated globular structures that possibly correspond to the glucan synthase complexes (Figure 16; Fèvre and Rougier, 1981).

Figure 16. Electron micrograph of glucan microfibrils synthesized in vitro by membrane preparations from S. monoica incubated in

absence of MgCl2 and in the presence of high substrate concentrations (1 mM) (× 70 000). The bound globular structures may correspond to trapped enzyme complexes (reprinted with permission from Fèvre and Rougier, 1981).

Attempts have been made to isolate the β‐(13) and β‐(14)‐glucan synthases from S. monoica, by combining detergent extraction of membrane‐bound proteins with subsequent purification by ultracentrifugation on glycerol gradients and product entrapment (Bulone et al., 1990). Product entrapment uses the fact that the enzyme is attached to its product after synthesis. By pelleting the insoluble product the enzyme is also enriched and partially purified. The enzymes could be separated into two fractions with proteins of 34, 48 and 50 kDa characteristic of the β‐(13)‐glucan synthase and proteins of 55‐60 kDa for the CesA activity. This has led to the identification of the 34 kDa‐protein as a β‐(13)‐glucan synthase activator that belongs to the annexin family (Bouzenzana et al., 2006). Recently, the enrichment of chitin synthase and β‐(13)‐glucan synthase in detergent‐resistant membranes of S. monoica was reported (Briolay et al., 2009). Interestingly no cellulose

40 synthase activity could be detected in the detergent‐resistant membranes. The lack of cellulose synthase activity in these membrane structures could be due to the absence of CesA catalytic subunits in the isolated detergent fractions. Another explanation could be that the cellulose synthase activity is lost during the preparation of the membrane structures due to an inactivation of the CesA by the detergent used for the extraction or because of a short halflife of the enzyme. It remains however to identify the catalytic subunits of the cellulose and β‐(13)‐glucan synthases in Oomycetes in the preparations used for in vitro synthesis of β‐glucans.

5.3 Chitin biosynthesis In this section, a summary of the current knowledge on chitin biosynthesis and chitin synthases is presented. As mentioned earlier, chitin occurs in many different organisms such as insects, shellfish and fungi. However, chitin synthesis has been most extensively studied in yeasts, insects and fungi. Here, the presentation is limited to chitin biosynthesis in yeasts, fungi (mostly genetic data) and Oomycetes.

Chs genes and classification in yeast and fungi The Chs genes can be divided into two divisions, which are further divided into seven classes based on structural similarities (Ruiz‐Herrera et al., 2002; Ruiz‐Herrera and Ortiz‐Castellanos, 2009; Lenardon et al., 2010a). Division 1 contains classes I‐III while division 2 contains classes IV‐ VII (Ruiz‐Herrera et al., 2002). In yeast, only classes I, II and IV are present while all classes of chitin synthases are represented in filamentous fungi (Ruiz‐Herrera and Ortiz‐Castellanos, 2009). All known Oomycete Chs cluster separately from the yeast and fungal Chs and belong to division 1, as opposed to all other non‐fungal Chs, which are clustered in division 2 (Ruiz‐ Herrera et al., 2002; Ruiz‐Herrera and Ortiz‐Castellanos, 2009). The focus in this summary is on yeast Chs from division I (classes I and II), since these are the closest to Oomycete Chs and also the most studied. Class IV from yeast will also be discussed briefly.

Function and regulation of yeast Chs from classes I, II and IV There are three Chs genes in the S. cerevisiae genome, ScChs1 (class I), ScChs2 (class II) and ScChs3 (class IV). All are expressed, but with different functions. ScChs1 is responsible for the major part of the chitin synthase activity measured in vitro and has been proposed to have a repair function during cytokinesis (Cabib et al., 1992; Roncero, 2002). However, a deletion in Chs1 in S. cerevisiae is not lethal, but cells are sensitive to acidic pH (Cabib et al., 1989). ScChs2 is crucial for primary septum formation and is therefore essential for the yeast (Shaw et al., 1991). ScChs3 is responsible for the major in vivo chitin synthase activity during cell wall formation and is also involved in bud scar formation (Shaw et al., 1991; Valdivieso et al., 1991). However, ScChs3 is not essential for the yeast, as opposed to Chs2. The zymogenic nature of Chs1, 2 and 3 and their regulation by partial proteolysis has been debated. In fact, it has been shown that Chs1, 2 and 3 are active without proteolytic digestion, but Chs1 and 2 activities are stimulated upon treatment with proteases whereas Chs3 is not (Cabib and Farkas, 1971; Sburlati and Cabib, 1986; Au‐Young and Robbins, 1990; Valdivieso et al., 1991; Uchida et al., 1996). Chitin biosynthesis takes place mostly in areas with active growth such as in the yeast bud tip or the fungal hyphal tip (Bowman and Free, 2006). Since chitin synthesis is important for cytokinesis and tip growth, the function of the various isoforms of Chs is tightly and differently regulated. Similar to the rapid turnover of cellulose synthase, the half‐life of ScChs2 is about 30 minutes in vivo, whereas ScChs3 has a half‐life of around 120 minutes (Chuang and Schekman, 1996). The S. cerevisiae Chs isoforms are also differently stimulated in vitro by divalent cations and they have different optimal pH (Choi and Cabib, 1994). Nickel ions have an inhibitory effect on ScChs1 and ScChs2 whereas ScChs3 is unaffected by this cation. Cobalt ions stimulate ScChs2 and ScChs3, but inhibit ScChs1. Proteolytic cleavage, as mentioned earlier, is another way of regulation used by the yeast. Results indicate that a proteolytically cleaved yeast activator, presumably a cysteine or serine protease, subsequently activates ScChs2 by proteolytic digestion (Martinez‐Rucobo et al., 2009). There are a number of results that support a phosphorylation‐dependent regulation of Chs. Mass spectromic analyses revealed that several phosphorylation sites occur in the N‐ terminus of the recombinantly expressed (E. coli and P. pastoris) and purified Chs2 from S. cerevisiae (Martinez‐Rucobo et al., 2009). A phosphatase treatment of the expressed recombinant ScChs2 indicates that the dephosphorylated protein is rapidly digested by

42 proteases. Contrarily, the phosphorylated form of ScChs2 or an N‐terminally truncated ScChs2 are stimulated by proteolytic digestion, pointing towards the role of phosphorylation in the regulation of the protein (Martinez‐Rucobo et al., 2009). Phosphorylation by cyclin‐ dependent kinase 1, a kinase involved in the control of the cell cycle, was also shown in vitro by using γ32P‐adenosine‐tri‐phosphate. In a phosphoproteomic study, several phosphorylation sites in the N‐terminus of Candida albicans Chs3 were identified (Lenardon et al., 2010b). An important serine was point mutated to alanine or glutamate to eliminate or mimic phosphorylation, respectively. Localisation studies with the mutant constructs fused to YFP in a Chs3 C. albicans deletion background showed impaired localisation of both the alanine and the glutamate mutants, indicating the importance of phosphorylation for the cellular localisation. The localisation was compared to that of a non‐mutated Chs3‐YFP fusion protein, which localised in the tip during cell growth and in the septum during cell division (Lenardon et al., 2007). Scaffolding and regulating proteins associated to the different Chs are also very important for the regulation of chitin synthesis, and have been studied mostly for Chs3. However, this falls beyond the scope of this thesis and is not discussed here (see for instance Lenardon (2010a) or Latge (2007) for a review related to this subject).

Approaches to study the function of expressed yeast chitin synthases The properties of membrane proteins make them technically challenging to study and, consequently, very few membrane‐bound glycosyltransferases have been successfully expressed, purified and characterized as recombinant proteins. This is especially true for cellulose synthases and FKS‐like proteins, for which the successful expression and characterization of a catalytically active recombinant protein has never been reported. Among chitin synthases, there are some examples of division 1 Chs that have been functionally investigated by complementation of Chs mutants, usually in S. cerevisiae (Au‐ Young and Robbins, 1990; Uchida et al., 1996). The classes I and II Chs are the most studied isoforms in vitro and complementation or overexpression of proteins from these classes have been achieved mostly in other fungi or yeast (Lenardon et al., 2010a). One example of a recombinantly expressed protein is Chs1 from S. cerevisiae (class I), which has been expressed in the yeast Schizosaccharomyces pombe and characterized already in the 80’s

(Bulawa et al., 1986). Members from class IV, like Chs3 from S. cerevisiae, have never been expressed heterologously in a functional form (Jimenez et al., 2010). Recently, the expression of the recombinant Chs2 from S. cerevisiae in the yeast P. pastoris and subsequent characterization of the protein was reported (Martinez‐Rucobo et al., 2009), where a partly purified solubilised protein extract was used for the phosphorylation studies mentioned in the previous section. However, Chs is like CesAs an unstable protein. This is illustrated by the lack of studies reporting a successful recombinant expression, purification and characterization of a Chs.

Chitin biosynthesis in Oomycetes Typically, up to two chitin synthase genes have been identified in the Oomycetes species in which chitin biosynthesis is studied. The Chs orthologues show more similarity than their paralogues (Mort‐Bontemps et al., 1997; Badreddine et al., 2008), but no general nomenclature taking into account similarities of sequence exists for these genes. In this thesis, we report the isolation and full‐length sequences of Chs1 and Chs2 from S. monoica. The full‐length of Chs1 was determined for the first time while our reported Chs 2 sequence differs from the previously published sequence (see article IV). In Phytophthora species, typically only one putative Chs gene has been found (Mort‐Bontemps et al., 1997; Tyler et al., 2006a; BLAST of P. infestans genome). The Oomycete S. monoica chitin synthases are active in vitro (Bulone et al., 1992; Briolay et al., 2009) and, like yeast division 1 Chs, chitin biosynthesis activity in S. monoica is activated by proteolysis. Chitin synthase activity has been shown to occur in detergent‐resistant membranes similar to lipid rafts in animal cells (Briolay et al., 2009). However, unlike β‐(13)‐glucan synthase, chitin synthase activity was not released from the detergent‐resistant membranes upon incubation with β‐ methylcyclodextrin, which interacts with sterols and disrupts protein/sterol interactions. Thus, even though the detergent‐resistant membranes were shown to be enriched in sterols, chitin synthase does not seem to be interacting directly with sterols.

5.4 Inhibitors of glycosyltransferase activities Some inhibitors that act on cell wall synthesis are used to inhibit microbial and plant growth, since the cell wall is vital for the microorganisms and plants. The cell wall composition and biosynthetic enzymes are different between plants and microorganisms, which sometimes can make cell wall formation a good and selective target for drugs. There are several compounds which exclusively inhibit cellulose biosynthesis or chitin synthesis, by acting directly on the catalytic subunit as competitive inhibitors or indirectly by inhibiting some other components of the biosynthetic pathways. Some cellulose synthase inhibitors have been used to study cellulose biosynthesis, for instance 2,6‐dichlorobenzonitrile (DCB), isoxaben, triazole carboxamides, quinclorac and thiazolidinone (reviewed in Sabba and Vaughn, 1999). For example, Arabidopsis mutants that exhibit resistance to the cellulose synthase inhibitors isoxaben and thiazolidinone have been analyzed and their resistance was correlated to modifications of the AtCesA3 gene (Scheible et al., 2001). The specific chitin synthase inhibitors nikkomycins and polyoxins have been used to investigate chitin biosynthesis, but they are rather poor antifungal drugs due to the limited cellular uptake of the drugs.

DCB DCB (Figure 17), is a specific but reversible inhibitor of cellulose biosynthesis in vivo at micromolar concentrations (Hogetsu et al., 1974; Sabba and Vaughn, 1999). However, it does not inhibit cellulose formation in vitro (Blaschek et al., 1985), which means that the addition of DCB to an in vitro synthesis reaction has no effect on the incorporation of glucose from UDP‐glucose into cellulose. DCB most likely acts in an indirect way, as it does not bind to the catalytic subunit of CesA (Delmer et al., 1987). DCB is affecting cell wall formation and cytokinesis but not nuclear division (Meyer and Hert, 1978). It specifically inhibits cellulose biosynthesis in plants (Montezinos and Delmer, 1980), which is accompanied by a reduced cellulose content in the cell walls of the treated plants (Shedletzky et al., 1990; Shedletzky et al., 1992; Melida et al., 2009). DCB causes accumulation of CesA proteins in tobacco cells and Arabidopsis seedlings, as well as of rosettes in the plasma membrane of cells from wheat roots (Herth, 1987; Nakagawa and Sakurai, 1998; DeBolt et al., 2007). A microtubule‐ associated protein, MAP20, is a secondary cell wall upregulated protein, which was shown to

45 bind DCB in vitro (Rajangam et al., 2008). In maize cell cultures habituated to DCB, which only differentiates primary walls, no MAP20 mRNA could be detected in the cells, supporting the role for MAP20 in secondary cell walls (Melida et al., 2010). To summarize, these observations altogether indicate that DCB, most likely indirectly, acts on protein regulation and transcription levels of genes involved in cell wall formation. The protist pathogen Acanthamoeba causes blinding keratitis as well as fatal granulomatous encephalitis (reviewed in Niederkorn et al., 1999). The inhibition of this pathogen is difficult as it easily switches to its cyst form, making it insensitive to anti‐ amoebic drugs. However, the cyst cell wall is composed of cellulose and it was reported that micromolar concentrations of DCB prevent encystment (Dudley et al., 2007), possibly by interfering with the cellulose biosynthesis machinery.

Figure 17. Structure of DCB.

Congo Red Congo Red (Figure 18) is a dye that interacts strongly with β‐glucans (Wood, 1980). The mycelium of S. monoica cultivated in the presence of Congo Red exhibits a reduced growth rate, an altered morphology and an increased incorporation of glucose into cell wall carbohydrates (Nodet et al., 1990 a). In addition, Congo Red is acting as an inhibitor in vitro of the β‐(14)‐glucan synthase activity at concentrations higher than 50 μg×ml‐1 and β‐(13)‐glucan synthase activity at lower concentrations (starting at 25 μgxml‐1; Nodet et al., 1990 b). Thus, Congo Red affects cellulose biosynthesis in Oomycetes, but its precise molecular mode of action in this process remains unclear.

Figure 18. Structure of Congo Red.

Polyoxins and Nikkomycins Polyoxins and nikkomycins are peptide nucleosides produced by the bacteria Streptomyces tendae and Streptomyces cacaoi, respectively. Structurally, they are analogues of UDP‐ GlcNAc (Figure 19) and act as competitive inhibitors of the chitin synthases. Polyoxins and nikkomycins are the best characterized inhibitors of chitin biosynthesis. In S. monoica, polyoxin D inhibits chitin synthase activity in vitro. However it only affects the mycelium growth slightly even though the chitin content is significantly decreased in the cell walls (Bulone et al., 1992).

Figure 19. Structure of UDP‐GlcNAc (A) and the chitin synthase inhibitors polyoxin D (B) and nikkomycin Z (C) (reprinted with permission from Hector, 1993). 47

6. PRESENT INVESTIGATION: AIM AND SUMMARY OF RESULTS

6.1 Aim of the present investigation As stressed at the beginning of the introduction, cellulose and chitin are the most abundant natural carbohydrates on earth. However, the current knowledge on the enzymes responsible for cellulose and chitin synthesis is very limited, concerning just a few model organisms. The functional studies are mostly based on indirect data. The limited amount of data can somewhat be explained by the membrane association of CesAs and Chs. As membrane proteins, with several transmembrane helices, they are problematic targets for biochemical characterization. The aim of this thesis is to make a contribution to the current knowledge on CesA and Chs gene characteristics and function. The main model systems chosen for the work presented here are different species of Oomycetes. These organisms represent an important economical problem in the aquaculture and agriculture industries worldwide. The focus of the project was to investigate the enzymes involved in cell wall biosynthesis, a vital process for Oomycetes, in the two model organisms P. infestans and S. monoica. These two species are representative of plant and animal pathogens, respectively. The composition of some Oomycete cell walls has been investigated to a great extent and it is one of the features that distinguish Oomycetes from fungi. However, the enzymes responsible for the biosynthesis of cell wall carbohydrates have hardly been investigated in Oomycetes, although they represent potential targets of inhibitors. In this thesis, the CesAs and Chs genes and their function have been investigated by gene expression profiling, in vivo and in vitro inhibitor studies and gene silencing combined with cell wall analysis and infection studies (paper I and II). The PH domain associated with some Oomycete CesAs has been expressed independently and functionally characterized (paper III). To understand the mechanism of an enzyme it is needed to study the protein in an isolated form. In this thesis, the results from the recombinant expression and functional characterization of Chs2 from S. monoica are presented (paper IV).

6.2 Materials and methods Materials and methods concerning the results summarized in this chapter are described in the related papers I‐IV. In addition to the data presented in the papers, we have identified the Saprolegnia parasitica CesA and Chs genes from the recently sequenced genome of this species. The genome of S. parasitica (www.broadinstitute.org) was analysed by BLAST (Altschul et al., 1997) using the SmCesA3 and SmCesA2 gene sequences. The BLAST search retrieved five full‐length predicted CesA genes, and one additional gene containing a predicted PH domain in the N‐terminus and a truncated Pfam 03552 cellulose synthase domain. The predicted sequence of the protein corresponding to the latter gene (SPRG_08607.2) was used in a protein BLAST search (http://www.ncbi.nlm.nih.gov/blast/Blast.cgi). The best four hits corresponded to CesA4 from Phytophthora species with an E value of 0.0 and scores of 879 to 873. If a guanine (G) is inserted at position 1884 of the putative SpCesA4 gene, a premature stop codon is removed and the open reading frame becomes full‐length and contains the full Pfam 03552 cellulose synthase domain. This nucleotide is present in SmCesA4, supporting a sequencing error to be the cause of its absence in the SpCesA4 genomic sequence identified from the draft genome. The conserved domains of the full‐length translated SpCesA sequences were analysed (http://www.ncbi.nlm.nih.gov/cdd/). The part of the genes corresponding to the Pfam 03552 domains of the predicted SpCesA genes, as well as the Oomycete CesA genes used in the phylogenetic tree of paper II, were aligned by MAFFT (Katoh et al., 2005). The alignment was subsequently checked for the best amino acid substitution model with Prottest (Abascal et al., 2005) and used for phylogenetic analysis with the PhyML 3.0 software (Guindon and Gascuel, 2003), based on the LG amino acid substitution model and a boot strap analysis with 100 replicates. The best tree topology search was chosen. A phylogenetic tree was made in MEGA version 5 (Kumar et al., 2008).

6.3 Results and discussion

6.3.1 Characterization of a novel group of CesA genes in Oomycetes (Papers I, II and unpublished results) Genomic data on Oomycetes have been available only since the end of 2006. In particular, an EST database and four Oomycete draft genomes have been published (Gajendran et al., 2006; Tyler et al., 2006a; Haas et al., 2009; Baxter et al., 2010). In 2009, the first draft of the S. parasitica genome was released (www.broadinstitute.org). The identification of the CesA genes from S. monoica was performed before the release of the S. parasitica genome. Therefore, the CesA genes in S. monoica were identified by sequence similarity searches of Oomycete EST databases and subsequent de novo cloning and sequencing (see Materials and Methods of paper II). A BLAST search in the Saprolegnia EST database with plant and bacterial CesA sequences allowed us to identify an EST sequence (ID = SPM13D7) that contained part of the putative processive glycosyltransferase motif (D, QXXRW). The most significant E values after BLAST analysis were of 6×10‐14 with the G. xylinus CesA sequence (accession number AB010645) and 10‐3 with AtCesA4 (accession number AAO15532). To amplify the full‐length gene corresponding to the identified EST from the species S. monoica (SmCesA), degenerated primers were designed and 3’ and 5’ RACE (rapid amplification of cDNA ends) were performed (Figure 20).

Figure 20. Principle of RACE: primers specific to internal parts of an EST sequence are used together with primers that match sequences ligated to the ends of the target mRNA sequence (5’cap and 3’ poly A tail). The mRNA ends are then amplified, cloned and sequenced, providing full‐length sequence information.

When the full‐length sequence of the first SmCesA gene was obtained, it was used to search the recently sequenced Phytophthora genomes for homologous gene models. Four gene models were found in each of the three sequenced genomes. These genes were named CesA1, CesA2, CesA3 and CesA4, and the SmCesA gene we identified first corresponded to CesA3 of Phytophthora species. In parallel to the bioinformatic work of paper II, peptide sequences from PiCesA1 were identified in a proteomic study of paper I. Since this was the first CesA identified in vivo, it was named CesA1, which is the reason for the nomenclature used in papers I and II. A Southern Blot analysis (Southern, 1975) was made using genomic DNA from S. monoica to determine the number of CesA homologues present in this species. According to the Southern Blot analysis, three homologues were present (Figure 1 in paper II). Degenerated primers were designed to amplify the full‐length sequences of the putative two other SmCesAs. RACE amplification of the sequence corresponding to SmCesA2 (full‐ length) and SmCesA4 (~30 % of full‐length) was achieved. The data of the Southern Blot analysis and RACE suggested that S. monoica does not have the CesA1 gene homologue

51 identified in the Phytophthora genomes. A phylogenetic analysis with MrBayes (Ronquist and Huelsenbeck, 2003) supported the hypothesis that the split of an ancestral gene into CesA1 and CesA2 in Phytophthora was a recent event which had not occurred in S. monoica (Figure 2 D in paper II). All the S. monoica and Phytophthora genes identified share the conserved motifs common to GT2 processive glycosyltransferases, i.e. two conserved D residues in domain A, one D residue and the QXXRW motif in domain B. An alignment of a part of the B domain is shown in Figure 1 A in paper II. Orthologous Oomycete CesA genes share a greater similarity to one another than paralogues do. The Oomycete CesA1, 2, 3 and 4 genes constitute a new group of genes in the CesA family, with some unique features (Figure 1 in paper I and Figure 2 B in paper II). In particular, the N‐terminal ends of CesA1, 2, and 4 from Phytophthora, and CesA2 and 4 from S. monoica, contain a PH domain (cd00821, http://www.ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi) believed to be involved in signaling, binding to lipids or attachment to membranes according to the Pfam database (http://pfam.sanger.ac.uk/). This is a remarkable feature since no other known CesA contains such domains. The Oomycete CesA3 genes do not contain a PH domain but they exhibit an extended N‐terminus with an extra cluster of seven predicted transmembrane helices (Figure 8). The genome of S. parasitica was made publicly available after the publication of paper II. Since S. parasitica is a closely related species to S. monoica, the identification of putative CesA genes in S. parasitica is relevant in this thesis. The genome was investigated as described in the materials and methods part. A total of five full‐length genes corresponding to CesA homologues were identified. After manual correction, one additional CesA gene corresponding to SpCesA4 was obtained. Two homologues of SmCesA3, named SpCesA3‐1 and SpCesA3‐2, were identified in S. parasitica. SpCesA3‐1 has a deletion in the C‐terminal end, causing the loss of one predicted transmembrane helix compared to the sequence of SmCesA3 and CesA3s from Phytophthora species. This truncation leads to a different predicted topology for SpCesA3, with a cytosolic N‐terminal domain and a non‐cytosolic C‐ terminal end. This predicted topology is unlikely, since both the C‐ and N‐ termini are predicted to be cytosolic in all other CesA3s. As seen in paper II (in the Materials and Methods part), incorrect predicted gene models can be a problem in draft genomes. The deletion in SpCesA3‐1 as well as the missing nucleotide in SpCesA4, might be due to incorrect gene model prediction, incorrect assembly of the contig or missing sequence information in

52 the assembly of the draft genome of S. parasitica. In Figure 21, a phylogenetic tree made with the Pfam 03552 cellulose synthase domain of the translated CesA sequences from Phytophthora and Saprolegnia species is shown. S. parasitica contain six CesA genes, as opposed to four CesA genes in the investigated Phytophthora species and three in S. monoica. Noticeably, S. parasitica has a unique duplication in the CesA3 gene. Moreover, one additional CesA gene containing a PH domain is found in S. parasitica. This additional gene is homologous to SmCesA2. Even in the species S. monoica with a lower number of CesA genes, cellulose synthesis is vital and is therefore a key process. Consequently, the higher number of genes found in S. parasitica might indicate functional redundancy of some of the CesA proteins, but this needs to be further investigated.

Figure 21. Cladogram of Oomycete CesAs including the six new putative genes from S. parasitica. The different subclasses are coloured in green (CesA1 and 2 from Phytophthora), yellow (CesA3s), red (CesA4s) and purple (CesA2‐like homologues from Saprolegnia). The tree was rooted with NpCesA from the cyanobacterium Nostoc punctiforme.

6.3.2 Expression pattern, localization and function of the CesAs from P. infestans (Paper I) The analysis of the expression levels of the putative CesAs from P. infestans was performed at different life cycle stages, by using quantitative Real‐Time PCR with SYBR Green (Materials and Methods of paper I). PiCesA3 was the dominantly expressed homologue in the mycelium

54 from P. infestans. PiCesA1 and PiCesA2 were up‐regulated during cyst and appressorium formation, but PiCesA3 was still the dominantly expressed gene (Figure 5 in paper I). PiCesA4 was expressed only in minute amounts in the mycelium, appressorium and cyst (Figure 5 in paper I). Immunolocalization studies in P. infestans showed that PiCesAs are localized in the growing tip and infection‐like vesicles of appressoria and cysts (Figure 9 in paper I). Antibodies produced against synthetic peptides corresponding to the amino acid sequence of SmCesA3 were used. The peptides were localized in the conserved CesA C‐terminal region, which allowed detection of PiCesAs. However, since a conserved region was used, the antibodies do not allow distinction of the different CesA homologues. The antibodies recognized CesA in purified plasma membrane fractions of P. infestans and S. monoica, but gave no positive signal in cytosolic fractions. This Western Blot analysis confirms the localization of the target CesA proteins in the plasma membranes of both species (Pages 2 and 3 of Supplementary material of paper I, which follows paper I in this thesis). Generation of gene knockouts is a significant technical challenge due to the diploidy of Oomycetes and stable transformation is of low efficiency. Thus, to circumvent these problems, the functional characterization of the PiCesA genes was performed using gene silencing by the RNA‐interference method established in P. infestans (Whisson et al., 2005). The gene encoding GFP (green fluorescent protein) was introduced in the p34GFN vector (Si‐ Ammour et al., 2003) and used as a non‐endogenous negative control. Lines of P. infestans were silenced for all four CesA homologues simultaneously, by transforming the strains with dsRNA. The target sequences are specific to each gene and correspond to the globular regions of the proteins. The expression levels of CesA genes in the transformed lines were analyzed by Real‐Time PCR to confirm gene silencing. Lines with different levels of silencing were obtained giving moderate to severe phenotypes. The phenotypes of the silenced appressoria were further investigated. Normally, at least 60 % of wild‐type P. infestans cysts produce appressoria in vitro. The number of appressoria produced in vitro was determined for each individual silenced line. The control lines with the p34GFN vector germinated and at least 60 % of the corresponding cysts formed normal appressoria, which indicates that the control lines behave as the wild‐type line. Control lines formed infection‐like vesicle structures while silenced lines did not. Some silenced lines produced either typical or aberrant appressoria, which ruptured at their tip, indicating a less resistant cell wall. Most of the individual lines had a clear reduction in the formation of normal appressoria. In addition,

55 a larger proportion of cysts produced germ tubes exhibiting abnormal appresorium‐like structures. The severity of the alteration of normal appresorium development and the number of aberrant appressoria produced correlated with the levels of the silencing of the CesA1‐4 genes in each silenced line (Figures 6 and 7 in paper I). The involvement of the PiCesA genes in cellulose synthesis in vivo was analyzed in a total of 22 lines. Appressoria corresponding to several of these lines were pooled to obtain three different groups as follows: six non‐silenced lines showing a normal phenotype formed group one, eight silenced lines with a severe phenotype represented group two and group three consisted of eight lines with a more moderate phenotype. The three pools were freeze‐dried and the cellulose extracted from the corresponding cell walls (see Materials and Methods in paper I). Enzymatic hydrolysis of the extracted cellulose was performed using an excess of a mixture of three recombinant specific cellulases from Thermofibida fusca expressed in E. coli (Cel6A, Cel6B and Cel9A; generous gift from Professor David B. Wilson, Cornell University, USA) to determine the cellulose content in the cell walls. An endo β‐ (13)‐glucanase from Trichoderma (Megazyme) was also used to detect the possible occurrence of β‐(13)‐glucans in the extracted material. Well‐characterized polysaccharides were used to verify enzyme specificity, as described in Materials and Methods and Supplementary Material of paper I, which follows paper I in the thesis. The kinetics of released reducing sugars was followed during 72 hours hydrolysis, using an assay adapted from Nelson‐Somogyi (Nelson, 1944). The kinetics of hydrolysis of the samples from the pooled lines exhibiting severe and moderate phenotypes reached a plateau after 72 hours, suggesting a complete hydrolysis of the total amount of cellulose originally present in the samples. In the case of the non‐silenced lines used as control, the release of reducing sugars continued to increase after 72 hours hydrolysis, indicating that hydrolysis in those samples had not reached completion (Figure 8 in paper I). The amount of reducing sugars released from the extracted wall material of the control line was approximately twice as high as in the samples corresponding to the silenced lines with severe and moderate phenotypes after 72 hours hydrolysis. Taken together, these results show that the cellulose content of the silenced lines is decreased compared to that of the control lines. Therefore, it can be concluded that at least some of the CesA1‐4 genes are involved in cellulose biosynthesis (Figure 8 in paper I).

Silencing of PiCesAs reduced the formation of appressoria and infection vesicle‐like structures, and was accompanied by the formation of aberrant appressorial structures. P. infestans requires a functional appressorium for a successful infection of potato. Thus, the effect of the cellulose synthesis inhibitor DCB was tested on the formation of appressoria and infection of potato. 40 μM and 100 μM of DCB had no significant effect on P. infestans growth, but the highest concentration of inhibitor resulted in a severe reduction of zoospore release and in the complete inhibition of cyst germination. At 40 μM DCB, a rupture of the appressoria was observed at the growing tip. In addition, the morphologies of the DCB‐ treated appressoria and germ tubes were identical to those observed for the CesA‐silenced lines (Figure 6 in paper I). The cell walls of the wild‐type were uniform when observed by transmission electron microscopy whereas the morphology and structures of the cell walls of the silenced and DCB‐treated lines were altered (Figure 11 in paper I). These results indicate that the treatment of P. infestans with DCB leads to similar appressorial and germ tube phenotypes as those observed for the PiCesAs silenced lines. Since cellulose synthesis is required for the formation of appressoria and normal germ tubes, it may also be needed for the infection process of the host tissues. Normally, zoospores of wild‐type P. infestans encyst and germinate upon contact with the host tissues. Germ tubes are usually short and produce swollen appressoria at their tips. Leaves are subsequently infected by penetration of the appressoria between epidermal cells or through stomata. An infection vesicle is then produced in the epidermal cell adjacent to the site of penetration. The need for cellulose biosynthesis in the infection process was tested by treating zoospores with 40 μM DCB and inoculating potato leaflets. The progress of the disease was recorded six days after inoculation. The DCB‐treated zoospores were found to be completely non‐pathogenic, whereas leaflets inoculated with untreated zoospores exhibited lesions and severe necrosis (Figure 12 E in paper I). Scanning electron microscopy observations 16 hours after inoculation showed that the control zoospores had encysted, produced many appressoria and appeared to have penetrated the host. The DCB treated zoospores had also encysted, but produced long germ tubes and aberrant appressoria‐like structures similar to those observed in vitro (Figure 12 A‐D in paper I).

6.3.3 Effect of cellulose synthesis inhibitors on mycelial growth of S. monoica, expression of CesA genes and glucan synthase activities (Paper II) A DCB treatment did not show any effect on mycelial growth in P. infestans, but the growth of S. monoica mycelium decreased in the presence of DCB and the mycelium layer on agar medium was thinner (Figure 22 and Figure 3 in paper II). Congo Red, an inhibitor of β‐glucan synthesis, also affected the growth of the mycelium of S. monoica, as reported earlier (Nodet et al., 1990 a) (Figure 23 and Figure 3 in paper II). Altogether, and based on the specificity of DCB, these data suggest that cellulose biosynthesis is more important for the mycelial growth of S. monoica than P. infestans.

A B

Figure 22. Growth of S. monoica mycelium in the presence of DCB (Machlis solid medium). A) Control plate containing 1% methanol (solvent of DCB). B) Plate containing 200 μM DCB in 1% methanol; the colony is smaller and thinner than in the control in Figure A.

A B

Figure 23. Growth of S. monoica mycelium in the presence of Congo Red (Machlis solid medium). A) Control plate without inhibitor. B) Plate containing 100 μM Congo Red; the colony is smaller than in the control in Figure A and exhibits a denser morphology.

The transcription levels of the SmCesA genes were investigated in the mycelium growing in the absence of inhibitor (Figure 24). The expression pattern of S. monoica CesAs in the mycelium follows that of P. infestans, with SmCesA3 being the dominantly expressed gene and SmCesA2 and 4 being expressed at much lower levels compared to SmCesA3.

Figure 24. SmCesA3 is the most highly expressed CesA gene in the mycelium of S. monoica. The analysis was made by quantitative Real‐Time PCR using SYBR Green, as described in paper II. The expression was normalised to the endogenous 18S RNA gene, which is constitutively expressed. The levels of expression were calibrated relative to the expression of SmCesA4 which was arbitrarily set to 1.

The effect of DCB and Congo Red on the transcription of the SmCesA genes was investigated by cultivating the mycelium of S. monoica in the presence of 200 μM DCB, 100 μM Congo Red, 1 % methanol or water. RNA was extracted at 0.5, 2, 4 and 8 hours after cultivation. The levels of expression of the SmCesA genes were analyzed by quantitative Real‐Time PCR (see Material and Methods in paper II) and normalized with respect to the constitutive expression of the 18S RNA gene. The relative levels of expression were obtained by dividing all expression values by the value from the corresponding control at 0.5 hour (Figure 25 A and B). The levels of up‐regulation of SmCesAs by DCB varied during the time course, with the highest effect (approximately 3 times up‐regulation) observed for SmCesA2 after 8 hours compared to the control sample at the same time point (Figure 25 A). Each CesA gene showed a decreased level of expression after 4 hours of growth in the presence of DCB or 1% methanol (Figure 25 A). This effect was somewhat reflected in the Congo Red treated samples (Figure 25 B). It might be due to transcriptional fluctuations during the growth of S. monoica. The SmCesA genes are up‐regulated by Congo Red with a maximum of 5 times increase in the level of expression for SmCesA3 after 8 hours growth (compared to the

60 control sample at the same time point, Figure 25 B). Taken together, these results indicate that cell wall biosynthesis or, alternatively, the cell wall structure, is disturbed by DCB and Congo Red, which leads to an increase in the transcriptional levels of SmCesAs.

A B

Figure 25. Effect of DCB and Congo Red on the expression levels of SmCesAs investigated by Real‐Time PCR. The figure is a reprint of Figure 4 in paper II. A) Expression levels of SmCesAs from the mycelium grown in the presence of DCB. A maximum of 3 times up‐ regulation can be seen for SmCesA2 after 8 hours. B) Expression levels of SmCesAs from the mycelium grown in the presence of Congo Red. A maximum of 5 times up‐regulation for SmCesA3 can be seen after 8 hours.

The effect of the inhibitors on β‐glucan synthesis was also tested by assaying enzymatic activities in membranes prepared from the mycelium of S. monoica grown in the presence of 200 μM DCB, 1 % methanol or 100 μM Congo Red. Total membranes were isolated after four days of cultivation and 500 μg protein was used in in vitro assays of glucan synthases. The incorporation of [14C]glucose from UDP‐[14C]glucose in ethanol‐insoluble β‐glucans was measured (Figures 5 A and B in paper II). Product characterization was performed using specific β‐glucanases (see Materials and Methods in paper II). The synthesized β‐glucans consisted of a mixture of β‐(13)‐glucan and cellulose (Figure 5 C in paper II). The amount of [14C]glucose incorporated was twice as high in the case of the enzyme extracted from the DCB‐treated mycelium than for to the control grown in the presence of 1 % methanol. The levels of activity were approximately 30 % higher for the Congo Red sample compared to the enzymatic sample from the untreated mycelium. Since DCB has no effect on in vitro synthesis of β‐glucans (Blaschek et al., 1985), these data suggest an increase in the amount of expressed glucan synthases (possibly CesAs) or in their specific activity. Congo Red is an inhibitor of glucan synthesis in vitro (Nodet et al., 1990 b). Carry over of Congo Red during the membrane extraction may have partially inhibited the in vitro synthesis of β‐glucans thereby leading to an underestimation of the actual levels of activity measured in vitro using enzymatic samples prepared from the Congo Red treated mycelium. These results are in good agreement with the observed increased expression of the SmCesA genes when S. monoica is grown in the presence of DCB or Congo Red.

6.3.4 Characterization of the PH domain from SmCesA2 (Paper III) The amino acid sequence of the PH domain from SmCesA2 (SmCesA2PH) was compared to characterized PH domains from human proteins in an alignment based on structure (see

Materials and Methods in paper III). The putative PtdIns(3,4)P2 and/or PtdIns(3,4,5)P3‐ binding motif (PPBM) is found in group I PH domains, located in the β‐sheet 1 and β‐sheet 2 and their connecting loop (see Figure 10 for the structure of a PH domain). Characterized PH domains containing this motif bind specifically and with high affinity to PtdIns(3,4)P2 and/or

PtdIns(3,4,5)P3. The alignment analysis showed that SmCesA2PH shares the PPBM motif with group I PH domains, indicating that it might bind to PtdIns(3,4)P2 and/or PtdIns(3,4,5)P3

(Figure 1 in paper III). SmCesA2PH was used in a BLAST search against proteins in the structure database (pdb structures, http://www.ncbi.nlm.nih.gov/blast/Blast.cgi). The C‐ terminal PH domain from the Tandem PH domain‐containing protein 1 (TAPP1; Figure 10 in section 5.1.3 Pleckstrin homology domains) was the best hit with an E value of 2e‐05. This protein binds specifically to PtdIns(3,4)P2 and co‐localizes with F‐actin in B‐lymphoma cells (Marshall et al., 2002). In order to characterize the SmCesA2PH experimentally, the domain was expressed as a histidine tagged protein in E. coli and purified by nickel ion‐immobilized metal‐affinity chromatography (IMAC) (Material and Methods in paper III). The identity of the purified protein was confirmed by MS analysis of the intact protein. The observed molecular mass of the expressed SmCesA2PH was identical to the expected molecular mass of 26.7 kDa (Figure 2 A and B in paper III). The purified SmCesA2PH was used in a protein‐lipid dot‐blot assay with different phosphoinositides and other types of phospholipids. According to the assay, SmCesA2PH binds preferentially PtdIns(3,4)P2 and PtdIns(3,4,5)P3, but also PtdIns(4,5)P2 and mono‐ phosphorylated PtdIns with a slightly lower affinity (Figure 3A in paper III). Some interactions could also be detected with phosphatidic acid, phosphatidylserine and PtdIns(3,5)P2, but to a lesser extent. To determine the binding preference of SmCesA2PH, an array protein‐lipid dot‐blot assay with varying concentrations of the different phosphorylated phosphoinositides was performed (Materials and Methods in paper III). The array protein‐ lipid assay indicated that SmCesA2PH binds to all phosphorylated phosphoinositides (Figure 3 B in paper III). Since some PH domains bind protein ligands, like F‐actin, the ability of SmCesA2PH to bind F‐actin and microtubules was tested in pull‐down experiments (Material and Methods in paper III). The pull‐down assays showed that SmCesA2PH binds to both F‐ actin and microtubules in vitro (Figure 4 A and B in paper III). To investigate the interactions of SmCesA2PH in vivo, the protein was expressed as a C‐terminally tagged emerald GFP (EmGFP) protein in human U2OS osteosarcoma cells (Materials and Methods in paper III). This system was chosen because of its simple and fast transfection method and the ease to determine protein localization in these cells, compared to for instance the tobacco leaf agroinfiltration expression system where a large vacuole complicates the interpretation of for instance membrane localization. In fixed U2OS cells, SmCesA2PH‐EmGFP was localized in the cytosol, nucleus, subnuclear compartments

(possibly corresponding to nucleoli, seen as bright dots in the nucleus) and in the distal borders of the cells (Figures 5 I, J, and M in paper IV). This localization was also seen in the cytosol localized control protein, chloramphenicol acetyltransferase (CAT) fused to EmGFP, except from the subnuclear compartment localizations (Figures 5 B and E in paper IV). CAT, expressed in the same cell type, showed an evenly dispersed localization in the cytoplasm and nucleus similar to that seen when expressing only EmGFP. The localization of SmCesA2PH to the subnuclear compartments might be caused by interaction with the nuclear PtdIns pool that exists in human cells. The cells were stained for F‐actin by fluorescent phalloidin. In the cells expressing SmCesA2PH‐EmGFP, co‐localisation of SmCesA2PH‐EmGFP with small actin filaments was seen in the distal borders of the cells, possibly corresponding to actin stress fiber ends in focal adhesion points. This was not seen in the cells expressing the control CAT‐EmGFP. Staining for microtubules was performed, but worked poorly and the results were not clear. To conclude, SmCesA2PH seems to be localized mainly in the cytosol and in the nucleus, where it is particularly targeted to subnuclear compartments which might contain PtdIns. SmCesA2PH can co‐localize with F‐ actin in vivo, as suggested earlier by the in vitro pull‐down assay results. The microtubule interaction in vivo still needs to be confirmed.

6.3.5 Characterization of the Chs genes in the Oomycetes S. monoica and S. parasitica (Paper IV and unpublished material) Two Chs genes from S. monoica genomic DNA, Chs1 and Chs2, have been cloned and sequenced previously (Mort‐Bontemps et al., 1997). However, only a fragment of Chs1 was obtained. In paper IV, the full‐length sequences of Chs1 and Chs2 were amplified by 5’ and 3’ RACE (Figure 20) from mycelial cDNA. The new sequences revealed a mis‐predicted stop codon in the 5’end of the previously reported genomic sequence of Chs1 (Mort‐Bontemps et al., 1997). The translated open reading frames of the new sequences contained a putative Pfam 04212 Microtubule Interacting and Trafficking (MIT) domain, present at the N‐terminus of both Chs1 and Chs2. The finding of this domain and the PH domain in the CesAs is in line with the genomic study that reports an increased number of novel multi‐domain proteins involved in primary metabolic pathways in Oomycetes (Morris et al., 2009). A possible function for the MIT domain could be in the targeting or regulation of the Chs. As for the

CesA genes, a Southern Blot analysis (Southern, 1975) was made with two Chs probes and three different restriction enzymes to determine the number of Chs homologues present in the genome of S. monoica. According to the Southern Blot analysis, only two homologues were present (Figure S1 in Supplementary material of paper IV, which follows paper IV in the thesis). A BLAST (Altschul et al., 1997) search of the S. parasitica genome revealed the presence of six Chs homologues, of which three contained the putative MIT domain. No other Oomycete genome has been reported to contain more than two chitin synthase genes, which suggests that chitin formation is central for S. parasitica considering that chitin is not present in all Oomycetes, or that there is functional redundancy among some of the Chs proteins.

6.3.6 In vitro chitin synthase activities of recombinant SmChs1 and SmChs2 (Paper IV) In order to characterize SmChs1 and SmChs2, the proteins were separately expressed as C‐ terminal eGFP fusion proteins in the yeast P. pastoris. The recombinant proteins were correctly targeted to the plasma membrane in the yeast cells. CHAPS‐solubilised membranes from yeast cells expressing the recombinant proteins were used in in vitro chitin synthase assays. The incorporation of [14C]GlcNAc from UDP‐[14C]GlcNAc into an ethanol‐insoluble product was measured (Figure 2 A in paper IV). The activity increased four times when the solubilised membrane fraction from the yeast cells expressing SmChs2 was used, compared to solubilised extracts from wild‐type yeast membranes or from yeast membranes expressing SmChs1. The latter enzyme was found to be inactive in this assay. The background activity observed in microsomes from the wild‐type yeast cells is caused by the Chs of P. pastoris. The chitin synthesized in vitro by SmChs2 was visualized by electron microscopy, which revealed the presence of chitin in the form of microcrystallites (Figure 2 B in paper IV). These crystallites were sensitive to hydrolysis by a specific chitinase from Serratia marcescens (see Materials and Methods in paper IV),. The solubilised membranes from wild‐type yeast cells or yeast cells expressing SmChs1 did not produce chitin microcrystallites. The synthesized microscrystallites and the radioactive in vitro product obtained from the recombinant SmChs2 were both sensitive to chitinase, which confirms that SmChs2 has chitin synthase activity. SmChs1 was not active in vitro, which might indicate that it has another role than synthesizing crystalline chitin in S. monoica.

To date, the effect of the chitin synthase inhibitor nikkomycin Z has not been tested in Oomycetes. The mycelium of S. monoica grown in potato dextrose agar plates in the presence of 50 µM nikkomycin Z for six hours contained about 50 % abnormal cells compared to the mycelium grown on control plates. When the concentration of inhibitor was increased to 200 µM, 80 % of the cells showed an aberrant phenotype. The abnormal cells exhibited swollen hyphae, and their tips burst, indicating an effect of nikkomycin Z on cell wall stability (Figure 3 C, D and E in paper IV). After 5 days, 50 % of the cells in the mycelium grown in the presence of 50 µM nikkomycin Z had survived, but all showed an abnormal phenotype. The chitin content in the corresponding cell walls was not significantly decreased, indicating that chitin deficiency was overcome. The mycelium grown in the presence of 500 µM nikkomycin Z was analysed by field‐emission scanning electron microscopy. There were very few cells growing in this condition and the mycelium seemed to crawl on the surface of the agar medium, avoiding contact with nikkomycin Z (Figure S5 C in Supplementary material of paper IV, which is shown after paper IV in the thesis). The incubation of S. parasitica in the presence of 200 µM nikkomycin Z almost abolished cell growth, indicating that S. parasitica is more sensitive to this drug than S. monoica. The effect of nikkomycin Z on the CHAPS‐solubilised recombinant SmChs2 expressed in P. pastoris was analysed. The apparent inhibition constant (Kiapp) of SmChs2 for nikkomycin Z was 4.6 ± 0.4 µM. The Km of the enzyme for UDP‐GlcNAc, was 150 ± 12 µM (Figure S4 in Supplementary material of paper IV). SmChs2 is competitively inhibited by nikkomycin Z. S. monoica cells are negatively affected, but the chitin content in their cell walls remains unaffected. This indicates a compensatory effect certainly due to an increased expression of the SmChs genes. The levels of expression of SmChs1 and SmChs2 were analysed by quantitative Real‐Time PCR using SYBR Green in the mycelium grown in the presence of 50 µM nikkomycin Z and in the absence of the inhibitor (Figure 4 in paper IV). The expression level of SmChs2 was found to be 100 000 times higher than that of SmChs1 in the cells grown in the absence of inhibitor, suggesting that SmChs2 is responsible for the bulk of the chitin synthesized in the cell wall, in line with the in vitro synthesis results. In the presence of inhibitor, the expression level of SmChs2 was increased by 70 %, supporting a compensatory effect with increased levels of SmChs2 in the cells activated in the presence of the drug. The expression level of SmChs1 was increased by 300 % in the presence of the

66 inhibitor. However, this still represents only a minimal fraction of the expression level of SmChs2 (Figure 4 in paper IV).

6.4 Conclusions and perspectives Here we report the isolation and characterization of the first Oomycete CesA genes in the plant pathogen P. infestans and the fish pathogen S. monoica. The small group of CesA genes found in Oomycetes is new in the CesA family. All the identified genes share the common motifs of CesAs from other organisms and other processive enzymes from the GT2 family. But in addition to the D,D,D and QXXRW motifs, some genes, namely CesA1,2 and 4 from Phytophthora and CesA2 and 4 from S. monoica and S. parasitica, also contain unique and novel features. The occurrence of N‐terminal PH domains is the most important distinguishing characteristic. In paper III, we have shown that the PH domain of SmCesA2 is similar to a human PH domain from the protein TAPP1. Like TAPP1, SmCesA2PH binds to

PtdIn(3,4)P2, but also to other phosphorylated PtdIns, with high affinity. Similar to TAPP1, SmCesA2PH can interact directly in vitro and can co‐localize in vivo with F‐actin. In addition, SmCesA2PH interacts with microtubules in vitro. The binding of a PH domain to both actin and microtubules has to our knowledge never been reported. The PH domain of Oomycete CesAs might be involved in targeting the CesAs to the growing tip, since we have shown that PiCesAs are localized in the tip during the growth of the germinating tube in P. infestans. F‐ actin is known to be enriched in the growing tip of some Oomycetes and could play a similar role to that of microtubules in plants, by guiding the CesA complex in the plasma membrane. The binding to microtubules and phosphorylated PtdIns could have a regulating function of the PH domain‐containing CesA1, 2 and 4. Cellulose biosynthesis is important for the infection life stages since the expression of CesA1, 2 and 4 is upregulated in P. Infestans during these stages. The CesA3 genes contain seven additional transmembrane helices in their N‐terminus, which represent a novel feature of Oomycete CesAs. The role of these additional transmembrane helices and the corresponding proteins need to be investigated in the future. The possibility of using cellulose and chitin biosynthesis as targets for the development of new Oomycete inhibitors seems promising. We have shown that cellulose biosynthesis in Oomycetes is important for infection in P. infestans. However, since there

67 seems to be no chitin in the cell wall of P. infestans, but the genome contains a Chs gene, putative chitin synthesis in P. infestans has to be analysed. The genome of S. parasitica contains numerous CesA and Chs genes, indicating that cell wall synthesis of these carbohydrates is important for this severe fish pathogen. S. parasitica was shown to be more sensitive than S. monoica to the chitin synthase inhibitor nikkomycin Z, which indicates that chitin biosynthesis is a promising target for tackling this very important pathogen. The role of S. parasitica CesAs and Chs genes in the infection process need to be further investigated. In summary, the results presented here highlight the potential of using CesAs and Chs as anti‐Oomycete drug targets. In this thesis, one membrane‐bound processive glycosyltransferase from family GT2 has been expressed as a functional recombinant GFP fusion protein in yeast, and was subsequently characterised. As presented in paper IV, we have directly demonstrated for the first time the activity of a chitin synthase from an Oomycete species. In conclusion, the biochemical studies presented here have provided new insights into the function of Chs and CesA proteins from different Oomycete species. Despite the progress made in our work on the identification and functional characterization of CesA and Chs genes, numerous questions remain to be addressed regarding cellulose and chitin biosynthesis in Oomycetes. For instance, it has not been demonstrated whether terminal complexes exist in Oomycetes and if they are organized as rosettes or linear complexes. The eventual mode of interaction of the different Oomycete CesA gene products is another open question, as well as the possible role of their different N‐terminal domains in the assembly of the cellulose synthase machinery. The involvement of specific Oomycete CesAs in specific life stages, as suggested by our preliminary expression analyses, needs to be further investigated for CesAs and Chs in S. parasitica. Similarly, the eventual expression of the Chs in P. infestans should be investigated. Even though certain interaction partners of the PH domain from CesA2 were identified in this work, the putative interaction between the PH domains from the CesAs 1, 2 and 4 should be examined, to investigate whether they are involved in the formation of the cellulose synthase machinery. This could be done by BiFC using the soluble PH domains or by combining the full‐length protein of one CesA isoform with a soluble PH domain from another CesA isoform in a membrane‐based yeast two‐hybrid assay. In our future work, we will focus on resolving some of these remaining issues.

7. ACKNOWLEDGEMENTS I slutet av min magisterutbildning gjorde jag olika projekt som en del av en forskarskola. KTH var det sista stoppet på min universitetstur, då jag testade Stockholms tre universitet. Det var tack vare professor Tuula Teeri som jag blev intresserad av att forska på KTH, på den tiden avdelningen för Träbioteknik. Dessförinnan hade jag varit mest intresserad av immunologi. Tuula gav mig chansen att förlänga stoppet till fem års doktorandstudier – vilket jag vill tacka henne för.

Efter något år av doktorandstudier och scoopade projekt blev vi en del av Biomime – och jag fick en ny handledare, nämligen professor Vincent Bulone. Thank you Vincent – I have learnt a lot from you. Nu blev jag introducerad till en helt ny värld jag aldrig tidigare bekymrat mig om – nämligen den som handlar om oomyceter. Under tidens gång bytte vi namn och blev Glykovetenskap, vilket passade mitt ”nya” ämne bättre. Så det som började i tron om att forska om växter och nya material slutade i en avhandling om potatismögel och andra patogena oomyceter.

Det är många som jag vill tacka för att ha hjälpt mig, samarbetat med mig eller på annat sätt bidragit till att den här avhandlingen blev slutförd. Tack till professor Harry Brumer för korrektionsläsning av avhandlingen och till Dr. Ines Ezcurra för experimentella tips. Christian Brown och Dr. Gea Guerriero ska ha ett särskilt tack för gott samarbete!

Om jag inte haft så bra sammarbetspartners utanför KTH hade nog den här avhandlingen låtit vänta på sig ett otal år till, så jag vill även tacka Dr. Pieter van West och hans medarbetare i Skottland, Dr. Lars Arvestad på SBC, Dr. Arne Lindqvist, min forna exjobbare Elvira Hukasova samt min ridkompis Dr. Katarina Gradin, alla på KI.

Jag hade aldrig kunnat ha så här kul under forskandets gång (trots alla misslyckade resultat, sena kvällar, korkade tabbar och annat) om jag inte haft mina kollegor, speciellt de som har delat rum med mig: Felicia, Erik, Dr. Gustav och Dr. Jens! Utöver dem vill jag särskilt lyfta fram mina forna kollegor Dr. Sofia Andersson (för alla luncher och after works), Dr. Nomchit Kaewthai (för inblick i den thailändska kulturen) och Dr. Linda Lindskog (för en bra introduktion och pep under resans gång). Jag vill även passa på att tacka alla som jobbar på 70 plan 2, nu och förr, och speciellt Fredrika, T.C., Oliver, Nuria, Ela, Ela, Sassa, Paula, Hanna och Markus.

Tack till gänget från forskarskolan (ni vet hur det är ☺) och till min gamla kursare Josipa! Ibland är det skönt att se att det finns ett liv utanför labbet med goda vännner, så tack Oda, Marie, Emelie, Hannah, Elin (min nyvunna mammakompis) samt alla andra mina vänner och bekanta.

Slutligen vill jag tacka de allra viktigaste – min familj! Tack mamma, pappa, Kaj och farmor för att ni funnits där. Isabelle, min syster, vill jag särskilt tacka för att du och jag uppskattar samma nörderier. Jag vill även tacka min släkt i Finland och min mans familj.

Under min doktorandtid har jag hunnit med en del av de stora sakerna i livet, som att gifta mig med min Daniel och få en liten dotter, underbara busärtan Siri. Jag är tacksam över att ni har varit tålmodiga med mina sena kvällar i labbet och stundals rätt stressiga perioder, och jag hoppas att vi får mycket tid framöver att njuta av livet!

8. REFERENCES

Abascal, F., Zardoya, R., and Posada, D. (2005). ProtTest: selection of best-fit models of protein evolution. Bioinformatics 21, 2104-2105. Agrios, G.N. (2005). Diseases caused by Oomycetes. In Plant pathology (Academic Press), pp. 409-428. Allam, A., and Marshall, A.J. (2005). Role of the adaptor proteins Bam32, TAPP1 and TAPP2 in lymphocyte activation. Immunol Lett 97, 7-17. Altschul, S.F., Madden, T.L., Schaffer, A.A., Zhang, J.H., Zhang, Z., Miller, W., and Lipman, D.J. (1997). Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25, 3389-3402. Anderson, I.J., Watkins, R.F., Samuelson, J., Spencer, D.F., Majoros, W.H., Gray, M.W., and Loftus, B.J. (2005). Gene discovery in the Acanthamoeba castellanii genome. Protist 156, 203-214. Andrivon, D. (1996). The origin of Phytophthora infestans populations present in Europe in the 1840s: a critical review of historical and scientific evidence. Plant Pathol 45, 1027- 1035. Arioli, T., Peng, L., Betzner, A.S., Burn, J., Wittke, W., Herth, W., Camilleri, C., Hofte, H., Plazinski, J., Birch, R., Cork, A., Glover, J., Redmond, J., and Williamson, R.E. (1998). Molecular analysis of cellulose biosynthesis in Arabidopsis. Science 279, 717-720. Aronson, J.M., and Lin, C.C. (1978). Hyphal wall chemistry of Leptomitus lacteus. Mycologia 70, 363-369. Atalla, R.H., and Vanderhart, D.L. (1984). Native cellulose: a composite of two distinct crystalline forms. Science 223, 283-285. Au-Young, J., and Robbins, P.W. (1990). Isolation of a chitin synthase gene (CHS1) from Candida albicans by expression in Saccharomyces cerevisiae. Mol Microbiol 4, 197- 207. Badreddine, I., Lafitte, C., Heux, L., Skandalis, N., Spanou, Z., Martinez, Y., Esquerre- Tugaye, M.T., Bulone, V., Dumas, B., and Bottin, A. (2008). Cell wall chitosaccharides are essential components and exposed patterns of the phytopathogenic Oomycete Aphanomyces euteiches. Eukaryot Cell 7, 1980-1993. Baldauf, S.L., Roger, A.J., Wenk-Siefert, I., and Doolittle, W.F. (2000). A kingdom-level phylogeny of eukaryotes based on combined protein data. Science 290, 972-977. Baraldi, E., Djinovic Carugo, K., Hyvonen, M., Surdo, P.L., Riley, A.M., Potter, B.V., O'Brien, R., Ladbury, J.E., and Saraste, M. (1999). Structure of the PH domain from Bruton's tyrosine kinase in complex with inositol 1,3,4,5-tetrakisphosphate. Structure 7, 449-460. Barratt, D.H.P., Derbyshire, P., Findlay, K., Pike, M., Wellner, N., Lunn, J., Feil, R., Simpson, C., Maule, A.J., and Smith, A.M. (2009). Normal growth of Arabidopsis requires cytosolic invertase but not sucrose synthase. Proc Natl Acad Sci U S A 106, 13124-13129. Bartnicki-Garcia, S. (1968). Cell wall chemistry, morphogenesis, and taxonomy of fungi. Annu Rev Microbiol 22, 87-108. Baskin, T.I., Betzner, A.S., Hoggart, R., Cork, A., and Williamson, R.E. (1992). Root morphology mutants in Arabidopsis thaliana. Aust J Plant Physiol 19, 427-437. Baxter, L., Tripathy, S., Ishaque, N., Boot, N., Cabral, A., Kemen, E., Thines, M., Ah- Fong, A., Anderson, R., Badejoko, W., Bittner-Eddy, P., Boore, J.L., Chibucos,

M.C., Coates, M., Dehal, P., Delehaunty, K., Dong, S., Downton, P., Dumas, B., Fabro, G., Fronick, C., Fuerstenberg, S.I., Fulton, L., Gaulin, E., Govers, F., Hughes, L., Humphray, S., Jiang, R.H., Judelson, H., Kamoun, S., Kyung, K., Meijer, H., Minx, P., Morris, P., Nelson, J., Phuntumart, V., Qutob, D., Rehmany, A., Rougon-Cardoso, A., Ryden, P., Torto-Alalibo, T., Studholme, D., Wang, Y., Win, J., Wood, J., Clifton, S.W., Rogers, J., Van den Ackerveken, G., Jones, J.D., McDowell, J.M., Beynon, J., and Tyler, B.M. (2010). Signatures of adaptation to obligate biotrophy in the Hyaloperonospora arabidopsidis genome. Science 330, 1549-1551. Bernal, A.J., Jensen, J.K., Harholt, J., Sorensen, S., Moller, I., Blaukopf, C., Johansen, B., de Lotto, R., Pauly, M., Scheller, H.V., and Willats, W.G. (2007). Disruption of AtCslD5 results in reduced growth, reduced xylan and homogalacturonan synthase activity and altered xylan occurrence in Arabidopsis. Plant J 52, 791-802. Blanton, R.L., Fuller, D., Iranfar, N., Grimson, M.J., and Loomis, W.F. (2000). The cellulose synthase gene of Dictyostelium. Proc Natl Acad Sci U S A 97, 2391-2396. Blaschek, W., Semler, U., and Franz, G. (1985). The influence of potential inhibitors on the in vivo and in vitro cell wall β-glucan biosynthesis in tobacco cells. J Plant Physiol 120, 457-470. Blum, M., Waldner, M., and Gisi, U. (2010a). A single point mutation in the novel PvCesA3 gene confers resistance to the carboxylic acid amide fungicide mandipropamid in Plasmopara viticola. Fungal Genet Biol 47, 499-510. Blum, M., Boehler, M., Randall, E., Young, V., Csukai, M., Kraus, S., Moulin, F., Scalliet, G., Avrova, A.O., Whisson, S.C., and Fonne-Pfister, R. (2010b). Mandipropamid targets the cellulose synthase-like PiCesA3 to inhibit cell wall biosynthesis in the Oomycete plant pathogen, Phytophthora infestans. Mol Plant Pathol 11, 227-243. Bodin, A., Backdahl, H., Risberg, B., and Gatenholm, P. (2007). Nano cellulose as a scaffold for tissue engineered blood vessels. Tissue Eng 13, 885-885. Bouzenzana, J., Pelosi, L., Briolay, A., Briolay, J., and Bulone, V. (2006). Identification of the first Oomycete annexin as a (1−>3)-β-D-glucan synthase activator. Mol Microbiol 62, 552-565. Bowman, S.M., and Free, S.J. (2006). The structure and synthesis of the fungal cell wall. Bioessays 28, 799-808. Briolay, A., Bouzenzana, J., Guichardant, M., Deshayes, C., Sindt, N., Bessueille, L., and Bulone, V. (2009). Cell wall polysaccharide synthases are located in detergent- resistant membrane microdomains in Oomycetes. Appl Environ Microbiol 75, 1938- 1949. Brown, E.D., Sewell, E.W.C., and Pereira, M.P. (2009). The wall teichoic acid polymerase TagF is non-processive in vitro and amenableto study using steady state kinetic analysis. J Biol Chem 284, 21132-21138. Brown, R.M. (1985). Cellulose microfibril assembly and orientation - recent developments. J Cell Sci, 13-32. Brown, R.M. (1996). The biosynthesis of cellulose. J Macromol Sci Pure A33, 1345-1373. Brownfield, L., Ford, K., Doblin, M.S., Newbigin, E., Read, S., and Bacic, A. (2007). Proteomic and biochemical evidence links the callose synthase in Nicotiana alata pollen tubes to the product of the NaGSL1 gene. Plant J 52, 147-156. Bulawa, C.E., Slater, M., Cabib, E., Auyoung, J., Sburlati, A., Adair, W.L., and Robbins, P.W. (1986). The Saccharomyces cerevisiae structural gene for chitin synthase is not required for chitin synthesis in vivo. Cell 46, 213-225.

Bulone, V., Girard, V., and Fèvre, M. (1990). Separation and partial purification of 1,3-β- glucan and 1,4-β-glucan synthases from Saprolegnia. Plant Physiol 94, 1748-1755. Bulone, V., Chanzy, H., Gay, L., Girard, V., and Fèvre, M. (1992). Characterization of chitin and chitin synthase from the cellulosic cell wall fungus Saprolegnia monoica. Exp Mycol 16, 8-21. Burton, R.A., Wilson, S.M., Hrmova, M., Harvey, A.J., Shirley, N.J., Medhurst, A., Stone, B.A., Newbigin, E.J., Bacic, A., and Fincher, G.B. (2006). Cellulose synthase-like CslF genes mediate the synthesis of cell wall (1,3;1,4)-β-D-glucans. Science 311, 1940-1942. Cabib, E., and Farkas, V. (1971). The control of morphogenesis: an enzymatic mechanism for the initiation of septum formation in yeast. Proc Natl Acad Sci U S A 68, 2052- 2056. Cabib, E., Silverman, S.J., and Shaw, J.A. (1992). Chitinase and chitin synthase 1: counterbalancing activities in cell separation of Saccharomyces cerevisiae. J Gen Microbiol 138, 97-102. Cabib, E., Sburlati, A., Bowers, B., and Silverman, S.J. (1989). Chitin synthase 1, an auxiliary enzyme for chitin synthesis in Saccharomyces cerevisiae. J Cell Biol 108, 1665-1672. Carpita, N.C., and Gibeaut, D.M. (1993). Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant J 3, 1-30. Chanzy, H., Imada, K., Mollard, A., Vuong, R., and Barnoud, F. (1979). Crystallographic aspects of sub-elementary cellulose fibrils occurring in the wall of rose cells cultured in vitro. Protoplasma 100, 303-316. Charnock, S.J., and Davies, G.J. (1999). Structure of the nucleotide-diphospho-sugar transferase, SpsA from Bacillus subtilis, in native and nucleotide-complexed forms. Biochemistry-Us 38, 6380-6385. Charnock, S.J., Henrissat, B., and Davies, G.J. (2001). Three-dimensional structures of UDP-sugar glycosyltransferases illuminate the biosynthesis of plant polysaccharides. J Plant Physiol 125, 527-531. Chen, S., Ehrhardt, D.W., and Somerville, C.R. (2010). Mutations of cellulose synthase (CESA1) phosphorylation sites modulate anisotropic cell expansion and bidirectional mobility of cellulose synthase. Proc Natl Acad Sci U S A 107, 17188-17193. Choi, W.J., and Cabib, E. (1994). The use of divalent cations and pH for the determination of specific yeast chitin synthetases. Anal Biochem 219, 368-372. Chuang, J.S., and Schekman, R.W. (1996). Differential trafficking and timed localization of two chitin synthase proteins, Chs2p and Chs3p. J Cell Biol 135, 597-610. Cocuron, J.C., Lerouxel, O., Drakakaki, G., Alonso, A.P., Liepman, A.H., Keegstra, K., Raikhel, N., and Wilkerson, C.G. (2007). A gene from the cellulose synthase-like C family encodes a β-1,4 glucan synthase. Proc Natl Acad Sci U S A 104, 8550-8555. Colombani, A., Djerbi, S., Bessueille, L., Blomqvist, K., Ohlsson, A., Berglund, T., Teeri, T.T., and Bulone, V. (2004). In vitro synthesis of (1→3)-β-D-glucan (callose) and cellulose by detergent extracts of membranes from cell suspension cultures of hybrid aspen. Cellulose 11, 313-327. Cosgrove, D.J. (2005). Growth of the plant cell wall. Nat Rev Mol Cell Biol 6, 850-861. Coutinho, P.M., and Henrissat, B. (1999). Carbohydrate-active enzymes: an integrated database approach. In Recent Advances in Carbohydrate Bioengineering (Cambridge: The Royal Society of Chemistry ), pp. 3-12. Coutinho, P.M., Deleury, E., Davies, G.J., and Henrissat, B. (2003). An evolving hierarchical family classification for glycosyltransferases. J Mol Biol 328, 307-317.

Crowell, E.F., Gonneau, M., Stierhof, Y.D., Höfte, H., and Vernhettes, S. (2010). Regulated trafficking of cellulose synthases. Curr Opin Plant Biol 13, 700-705. Crowell, E.F., Bischoff, V., Desprez, T., Rolland, A., Stierhof, Y.D., Schumacher, K., Gonneau, M., Höfte, H., and Vernhettes, S. (2009). Pausing of Golgi bodies on microtubules regulates secretion of cellulose synthase complexes in Arabidopsis. Plant Cell 21, 1141-1154. Cui, X.J., Shin, H.S., Song, C., Laosinchai, W., Amano, Y., and Brown, R.M. (2001). A putative plant homolog of the yeast β-1,3-glucan synthase subunit FKS1 from cotton (Gossypium hirsutum L.) fibers. Planta 213, 223-230. Davies, G., and Henrissat, B. (1995). Structures and mechanisms of glycosyl hydrolases. Structure 3, 853-859. De Schutter, K., Lin, Y.C., Tiels, P., Van Hecke, A., Glinka, S., Weber-Lehmann, J., Rouze, P., Van de Peer, Y., and Callewaert, N. (2009). Genome sequence of the recombinant protein production host Pichia pastoris. Nat Biotechnol 27, 561-566. DeAngelis, P.L. (1999). Molecular directionality of polysaccharide polymerization by the Pasteurella multocida hyaluronan synthase. J Biol Chem 274, 26557-26562. DeBolt, S., Gutierrez, R., Ehrhardt, D.W., and Somerville, C. (2007). Nonmotile cellulose synthase subunits repeatedly accumulate within localized regions at the plasma membrane in Arabidopsis hypocotyl cells following 2,6-dichlorobenzonitrile treatment. Plant Physiol 145, 334-338. DeBolt, S., Scheible, W.R., Schrick, K., Auer, M., Beisson, F., Bischoff, V., Bouvier- Nave, P., Carroll, A., Hematy, K., Li, Y.H., Milne, J., Nair, M., Schaller, H., Zemla, M., and Somerville, C. (2009). Mutations in UDP-glucose:sterol glucosyltransferase in Arabidopsis cause transparent testa phenotype and suberization defect in seeds. Plant Physiol 151, 78-87. Delmer, D.P. (1999). Cellulose Biosynthesis: Exciting times for a difficult field of study. Annu Rev Plant Physiol Plant Mol Biol 50, 245-276. Delmer, D.P., Read, S.M., and Cooper, G. (1987). Identification of a receptor protein in cotton fibers for the herbicide 2,6-dichlorobenzonitrile. Plant Physiol 84, 415-420. Desprez, T., Juraniec, M., Crowell, E.F., Jouy, H., Pochylova, Z., Parcy, F., Höfte, H., Gonneau, M., and Vernhettes, S. (2007). Organization of cellulose synthase complexes involved in primary cell wall synthesis in Arabidopsis thaliana. Proc Natl Acad Sci U S A 104, 15572-15577. Dhe-Paganon, S., Ottinger, E.A., Nolte, R.T., Eck, M.J., and Shoelson, S.E. (1999). Crystal structure of the pleckstrin homology-phosphotyrosine binding (PH-PTB) targeting region of insulin receptor substrate 1. Proc Natl Acad Sci U S A 96, 8378- 8383. Dhugga, K.S., Barreiro, R., Whitten, B., Stecca, K., Hazebroek, J., Randhawa, G.S., Dolan, M., Kinney, A.J., Tomes, D., Nichols, S., and Anderson, P. (2004). Guar seed β-mannan synthase is a member of the cellulose synthase super gene family. Science 303, 363-366. Djerbi, S., Lindskog, M., Arvestad, L., Sterky, F., and Teeri, T.T. (2005). The genome sequence of black cottonwood (Populus trichocarpa) reveals 18 conserved cellulose synthase (CesA) genes. Planta 221, 739-746. Djerbi, S., Aspeborg, H., Nilsson, P., Sundberg, B., Mellerowicz, E., Blomqvist, K., and Teeri, T.T. (2004). Identification and expression analysis of genes encoding putative cellulose synthases (CesA) in the hybrid aspen, Populus tremula (L.) x P. tremuloides (Michx.). Cellulose 11, 301-312. Doblin, M.S., Pettolino, F.A., Wilson, S.M., Campbell, R., Burton, R.A., Fincher, G.B., Newbigin, E., and Bacic, A. (2009). A barley cellulose synthase-like CslH gene

mediates (1,3;1,4)-β-D-glucan synthesis in transgenic Arabidopsis. Proc Natl Acad Sci U S A 106, 5996-6001. Douglas, C.M., Foor, F., Marrinan, J.A., Morin, N., Nielsen, J.B., Dahl, A.M., Mazur, P., Baginsky, W., Li, W., el-Sherbeini, M., and et al. (1994). The Saccharomyces cerevisiae FKS1 (ETG1) gene encodes an integral membrane protein which is a subunit of 1,3-β-D-glucan synthase. Proc Natl Acad Sci U S A 91, 12907-12911. Dowler, S., Currie, R.A., Downes, C.P., and Alessi, D.R. (1999). DAPP1: a dual adaptor for phosphotyrosine and 3-phosphoinositides. Biochem J 342 ( Pt 1), 7-12. Downing, A.K., Driscoll, P.C., Gout, I., Salim, K., Zvelebil, M.J., and Waterfield, M.D. (1994). Three-dimensional solution structure of the pleckstrin homology domain from dynamin. Curr Biol 4, 884-891. Dudley, R., Alsam, S., and Khan, N.A. (2007). Cellulose biosynthesis pathway is a potential target in the improved treatment of Acanthamoeba keratitis. Appl Microbiol Biotechnol 75, 133-140. Dutta, P.K., Dutta, J., and Tripathi, V.S. (2004). Chitin and chitosan: Chemistry, properties and applications. J Sci Industr Res 63, 20-31. Endler, A., and Persson, S. (2011). Cellulose synthases and synthesis in Arabidopsis. Mol Plant 4, 199-211. Ferguson, K.M., Lemmon, M.A., Schlessinger, J., and Sigler, P.B. (1995). Structure of the high affinity complex of inositol trisphosphate with a phospholipase C pleckstrin homology domain. Cell 83, 1037-1046. Fèvre, M., and Rougier, M. (1981). β-1-3-glucan and β-1-4-glucan synthesis by membrane- fractions from the fungus Saprolegnia. Planta 151, 232-241. Fry, W.E., and Goodwin, S.B. (1997). Re-emergence of potato and tomato late blight in the United States. Plant Dis 81, 1349-1357. Fugelstad, J., Bouzenzana, J., Djerbi, S., Guerriero, G., Ezcurra, I., Teeri, T.T., Arvestad, L., and Bulone, V. (2009). Identification of the cellulose synthase genes from the Oomycete Saprolegnia monoica and effect of cellulose synthesis inhibitors on gene expression and enzyme activity. Fungal Genet Biol 46, 759-767. Fujii, S., Hayashi, T., and Mizuno, K. (2010). Sucrose synthase is an integral component of the cellulose synthesis machinery. Plant Cell Physiol 51, 294-301. Fushman, D., Najmabadi-Haske, T., Cahill, S., Zheng, J., LeVine, H., 3rd, and Cowburn, D. (1998). The solution structure and dynamics of the pleckstrin homology domain of G protein-coupled receptor kinase 2 (β-adrenergic receptor kinase 1). A binding partner of Gβγ subunits. J Biol Chem 273, 2835-2843. Gajendran, K., Gonzales, M.D., Farmer, A., Archuleta, E., Win, J., Waugh, M.E., and Kamoun, S. (2006). Phytophthora functional genomics database (PFGD): functional genomics of Phytophthora-plant interactions. Nucleic Acids Res 34, D465-470. Glaser, L. (1958). The synthesis of cellulose in cell-free extracts of Acetobacter xylinum. J Biol Chem 232, 627-636. Goubet, F., Barton, C.J., Mortimer, J.C., Yu, X., Zhang, Z., Miles, G.P., Richens, J., Liepman, A.H., Seffen, K., and Dupree, P. (2009). Cell wall glucomannan in Arabidopsis is synthesised by CslA glycosyltransferases, and influences the progression of embryogenesis. Plant J 60, 527-538. Gray, A., Van Der Kaay, J., and Downes, C.P. (1999). The pleckstrin homology domains of protein kinase B and GRP1 (general receptor for phosphoinositides-1) are sensitive and selective probes for the cellular detection of phosphatidylinositol 3,4-bisphosphate and/or phosphatidylinositol 3,4,5-trisphosphate in vivo. Biochem. J. 344, 929-936. Grenville-Briggs, L.J., and van West, P. (2005). The biotrophic stages of Oomycete-plant interactions. Adv Appl Microbiol 57, 217-243.

Grenville-Briggs, L.J., Anderson, V.L., Fugelstad, J., Avrova, A.O., Bouzenzana, J., Williams, A., Wawra, S., Whisson, S.C., Birch, P.R., Bulone, V., and van West, P. (2008). Cellulose synthesis in Phytophthora infestans is required for normal appressorium formation and successful infection of potato. Plant Cell 20, 720-738. Guerriero, G., Fugelstad, J., and Bulone, V. (2010). What do we really know about cellulose biosynthesis in higher plants? J Integr Plant Biol 52, 161-175. Guindon, S., and Gascuel, O. (2003). A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol 52, 696-704. Gutierrez, R., Lindeboom, J.J., Paredez, A.R., Emons, A.M., and Ehrhardt, D.W. (2009). Arabidopsis cortical microtubules position cellulose synthase delivery to the plasma membrane and interact with cellulose synthase trafficking compartments. Nat Cell Biol 11, 797-806. Haas, B.J., Kamoun, S., Zody, M.C., Jiang, R.H., Handsaker, R.E., Cano, L.M., Grabherr, M., Kodira, C.D., Raffaele, S., Torto-Alalibo, T., Bozkurt, T.O., Ah- Fong, A.M., Alvarado, L., Anderson, V.L., Armstrong, M.R., Avrova, A., Baxter, L., Beynon, J., Boevink, P.C., Bollmann, S.R., Bos, J.I., Bulone, V., Cai, G., Cakir, C., Carrington, J.C., Chawner, M., Conti, L., Costanzo, S., Ewan, R., Fahlgren, N., Fischbach, M.A., Fugelstad, J., Gilroy, E.M., Gnerre, S., Green, P.J., Grenville-Briggs, L.J., Griffith, J., Grunwald, N.J., Horn, K., Horner, N.R., Hu, C.H., Huitema, E., Jeong, D.H., Jones, A.M., Jones, J.D., Jones, R.W., Karlsson, E.K., Kunjeti, S.G., Lamour, K., Liu, Z., Ma, L., Maclean, D., Chibucos, M.C., McDonald, H., McWalters, J., Meijer, H.J., Morgan, W., Morris, P.F., Munro, C.A., O'Neill, K., Ospina-Giraldo, M., Pinzon, A., Pritchard, L., Ramsahoye, B., Ren, Q., Restrepo, S., Roy, S., Sadanandom, A., Savidor, A., Schornack, S., Schwartz, D.C., Schumann, U.D., Schwessinger, B., Seyer, L., Sharpe, T., Silvar, C., Song, J., Studholme, D.J., Sykes, S., Thines, M., van de Vondervoort, P.J., Phuntumart, V., Wawra, S., Weide, R., Win, J., Young, C., Zhou, S., Fry, W., Meyers, B.C., van West, P., Ristaino, J., Govers, F., Birch, P.R., Whisson, S.C., Judelson, H.S., and Nusbaum, C. (2009). Genome sequence and analysis of the Irish potato famine pathogen Phytophthora infestans. Nature 461, 393-398. Han, N.S., and Robyt, J.F. (1998). The mechanism of Acetobacter xylinum cellulose biosynthesis: direction of chain elongation and the role of lipid pyrophosphate intermediates in the cell membrane. Carbohydr Res 313, 125-133. Hardham, A.R. (2007). Cell biology of plant-Oomycete interactions. Cell Microbiol 9, 31- 39. Haslam, R.J., Koide, H.B., and Hemmings, B.A. (1993). Pleckstrin domain homology. Nature 363, 309-310. Hector, R.F. (1993). Compounds active against cell-walls of medically important fungi. Clin Microbiol Rev 6, 1-21. Helbert, W., Sugiyama, J., Ishihara, M., and Yamanaka, S. (1997). Characterization of native crystalline cellulose in the cell walls of Oomycota. J Biotechnol 57, 29-37. Hematy, K., Sado, P.E., Van Tuinen, A., Rochange, S., Desnos, T., Balzergue, S., Pelletier, S., Renou, J.P., and Hofte, H. (2007). A receptor-like kinase mediates the response of Arabidopsis cells to the inhibition of cellulose synthesis. Curr Biol 17, 922-931. Henrissat, B., Coutinho, P.M., and Davies, G.J. (2001). A census of carbohydrate-active enzymes in the genome of Arabidopsis thaliana. Plant Mol Biol 47, 55-72. Herth, W. (1987). Effects of 2,6-DCB on plasma-membrane rosettes of wheat root cells. Naturwissenschaften 74, 556-557.

Hirano, S. (1999). Chitin and chitosan as novel biotechnological materials. Pol International 48, 732-734. Hogetsu, T., Shibaoka, H., and Shimokoriyama, M. (1974). Involvement of cellulose synthesis in actions of gibberellin and kinetin on cell expansion. 2,6- dicholorobenzonitrile as a new cellulose synthesis inhibitor. Plant Cell Physiol 15, 389-393. Hong, Z.L., Delauney, A.J., and Verma, D.P.S. (2001). A cell plate specific callose synthase and its interaction with phragmoplastin. Plant Cell 13, 755-768. Hu, S., Gao, Y., Tajima, K., Yao, M., Yoda, T., Shimura, D., Satoh, Y., Kawano, S., Tanaka, I., and Munekata, M. (2008). Purification, crystallization and preliminary X-ray studies of AxCesD required for efficient cellulose biosynthesis in Acetobacter xylinum. Prot Pept Lett 15, 115-117. Hu, S.Q., Gao, Y.G., Tajima, K., Sunagawa, N., Zhou, Y., Kawano, S., Fujiwara, T., Yoda, T., Shimura, D., Satoh, Y., Munekata, M., Tanaka, I., and Yao, M. (2010). Structure of bacterial cellulose synthase subunit D octamer with four inner passageways. Proc Natl Acad Sci U S A 107, 17957-17961. Hudson, S.M., and Smith, C. (1998). The structure and compostition of chitin and chitosan. In Biopolymers from renewable resources, D.L. Kaplan, ed (Berlin Heidelberg: Springer-Verlag), pp. 101. Hyvonen, M., and Saraste, M. (1997). Structure of the PH domain and Btk motif from Bruton's tyrosine kinase: molecular explanations for X-linked agammaglobulinaemia. EMBO J 16, 3396-3404. Imai, T., Watanabe, T., Yui, T., and Sugiyama, J. (2003). The directionality of chitin biosynthesis: a revisit. Biochem J 374, 755-760. Isakoff, S.J., Cardozo, T., Andreev, J., Li, Z., Ferguson, K.M., Abagyan, R., Lemmon, M.A., Aronheim, A., and Skolnik, E.Y. (1998). Identification and analysis of PH domain-containing targets of phosphatidylinositol 3-kinase using a novel in vivo assay in yeast. EMBO J 17, 5374-5387. Jacob-Wilk, D., Kurek, I., Hogan, P., and Delmer, D.P. (2006). The cotton fiber zinc- binding domain of cellulose synthase A1 from Gossypium hirsutum displays rapid turnover in vitro and in vivo. Proc Natl Acad Sci U S A 103, 12191-12196. Jarvis, M. (2003). Chemistry: cellulose stacks up. Nature 426, 611-612. Jayakumar, R., Nwe, N., Tokura, S., and Tamura, H. (2007). Sulfated chitin and chitosan as novel biomaterials. Int J Biol Macromol 40, 175-181. Jimenez, C., Sacristan, C., Roncero, M.I., and Roncero, C. (2010). Amino acid divergence between the Chs domain contributes to the different intracellular behaviour of Family II fungal chitin synthases in Saccharomyces cerevisiae. Fungal Genet Biol 47, 1034- 1043. Joshi, C.P., Thammannagowda, S., Fujino, T., Gou, J.Q., Avci, U., Haigler, C.H., McDonnell, L.M., Mansfield, S.D., Mengesha, B., Carpita, N.C., Harris, D., Debolt, S., and Peter, G.F. (2011). Perturbation of wood cellulose synthesis causes pleiotropic effects in transgenic aspen. Mol Plant 4, 331-345. Judelson, H.S., and Ah-Fong, A.M. (2010). The kinome of Phytophthora infestans reveals Oomycete-specific innovations and links to other taxonomic groups. BMC Genomics 11, 700. Kang, M.S., and Cabib, E. (1986). Regulation of fungal cell-wall growth - a guanine nucleotide-binding,proteinaceous component required for activity of (1-->3)-β-D- glucan synthase. Proc Natl Acad Sci U S A 83, 5808-5812. Katoh, K., Kuma, K., Toh, H., and Miyata, T. (2005). MAFFT version 5: improvement in accuracy of multiple sequence alignment. Nucleic Acids Res 33, 511-518.

Kavran, J.M., Klein, D.E., Lee, A., Falasca, M., Isakoff, S.J., Skolnik, E.Y., and Lemmon, M.A. (1998). Specificity and promiscuity in phosphoinositide binding by pleckstrin homology domains. J Biol Chem 273, 30497-30508. Keegstra, K. (2010). Plant cell walls. Plant Physiol 154, 483-486. Kimura, S., Laosinchai, W., Itoh, T., Cui, X.J., Linder, C.R., and Brown, R.M. (1999). Immunogold labeling of rosette terminal cellulose synthesizing complexes in the vascular plant Vigna angularis. Plant Cell 11, 2075-2085. Klemm, D., Heublein, B., Fink, H.P., and Bohn, A. (2005). Cellulose: Fascinating biopolymer and sustainable raw material. Angew Chem Int Edit 44, 3358-3393. Koshiba, S., Kigawa, T., Kim, J.H., Shirouzu, M., Bowtell, D., and Yokoyama, S. (1997). The solution structure of the pleckstrin homology domain of mouse Son-of-sevenless 1 (mSos1). J Mol Biol 269, 579-591. Koyama, M., Helbert, W., Imai, T., Sugiyama, J., and Henrissat, B. (1997). Parallel-up structure evidences the molecular directionality during biosynthesis of bacterial cellulose. Proc Natl Acad Sci U S A 94, 9091-9095. Kudlicka, K., Lee, J.H., and Brown, R.M. (1996). A comparative analysis of in vitro cellulose synthesis from cell-free extracts of mung bean (Vigna radiata, Fabaceae) and cotton (Gossypium hirsutum, Malvaceae). Am J Bot 83, 274-284. Kumar, S., and Rzhetsky, A. (1996). Evolutionary relationships of eukaryotic kingdoms. J Mol Evol 42, 183-193. Kumar, S., Nei, M., Dudley, J., and Tamura, K. (2008). MEGA: a biologist-centric software for evolutionary analysis of DNA and protein sequences. Brief Bioinform 9, 299-306. Kurek, I., Kawagoe, Y., Jacob-Wilk, D., Doblin, M., and Delmer, D. (2002). Dimerization of cotton fiber cellulose synthase catalytic subunits occurs via oxidation of the zinc- binding domains. Proc Natl Acad Sci U S A 99, 11109-11114. Kurita, K. (2006). Chitin and chitosan: functional biopolymers from marine crustaceans. Mar Biotechnol (NY) 8, 203-226. Lai-Kee-Him, J., Chanzy, H., Muller, M., Putaux, J.L., Imai, T., and Bulone, V. (2002). In vitro versus in vivo cellulose microfibrils from plant primary wall synthases: Structural differences. J Biol Chem 277, 36931-36939. Lairson, L.L., Henrissat, B., Davies, G.J., and Withers, S.G. (2008). Glycosyltransferases: Structures, functions and mechanisms. Annu Rev Biochem 77, 25.21-25.35. Lartseva, L.V. (1986). Saprolegniaceae on the spawn of sturgeons and salmons fishes. Hydrobiol J 22, 103–107. Latge, J.P. (2007). The cell wall: a carbohydrate armour for the fungal cell. Mol Microbiol 66, 279-290. Leevers, S.J., Vanhaesebroeck, B., and Waterfield, M.D. (1999). Signalling through phosphoinositide 3-kinases: the lipids take centre stage. Curr Opin Cell Biol 11, 219- 225. Lemmon, M.A. (2004). Pleckstrin homology domains: not just for phosphoinositides. Biochem Soc T 32, 707-711. Lemmon, M.A. (2008). Membrane recognition by phospholipid-binding domains. Nat Rev Mol Cell Biol 9, 99-111. Lemmon, M.A., and Ferguson, K.M. (2000). Signal-dependent membrane targeting by pleckstrin homology (PH) domains. Biochem J 350, 1-18. Lemmon, M.A., Ferguson, K.M., O'Brien, R., Sigler, P.B., and Schlessinger, J. (1995). Specific and high-affinity binding of inositol phosphates to an isolated pleckstrin homology domain. Proc Natl Acad Sci U S A 92, 10472-10476.

Lenardon, M.D., Munro, C.A., and Gow, N.A.R. (2010a). Chitin synthesis and fungal pathogenesis. Curr Opin Microbiol 13, 416-423. Lenardon, M.D., Whitton, R.K., Munro, C.A., Marshall, D., and Gow, N.A. (2007). Individual chitin synthase enzymes synthesize microfibrils of differing structure at specific locations in the Candida albicans cell wall. Mol Microbiol 66, 1164-1173. Lenardon, M.D., Milne, S.A., Mora-Montes, H.M., Kaffarnik, F.A.R., Peck, S.C., Brown, A.J.P., Munro, C.A., and Gow, N.A.R. (2010b). Phosphorylation regulates polarisation of chitin synthesis in Candida albicans. J Cell Sci 123, 2199-2206. Li, J., Burton, R.A., Harvey, A.J., Hrmova, M., Wardak, A.Z., Stone, B.A., and Fincher, G.B. (2003). Biochemical evidence linking a putative callose synthase gene with (1 -- >3)-β-D-glucan biosynthesis in barley. Plant Mol Biol 53, 213-225. Liepman, A.H., Wilkerson, C.G., and Keegstra, K. (2005). Expression of cellulose synthase-like (Csl) genes in insect cells reveals that CslA family members encode mannan synthases. Proc Natl Acad Sci U S A 102, 2221-2226. Lin, C.C., and Aronson, J.M. (1970). Chitin and cellulose in the cell walls of the Oomycete, Apodachlya sp. Arch Mikrobiol 72, 111-114. Lin, F.C., Brown, R.M., Jr., Drake, R.R., Jr., and Haley, B.E. (1990). Identification of the uridine 5'-diphosphoglucose (UDP-Glc) binding subunit of cellulose synthase in Acetobacter xylinum using the photoaffinity probe 5-azido-UDP-Glc. J Biol Chem 265, 4782-4784. Lloyd, C., and Chan, J. (2008). The parallel lives of microtubules and cellulose microfibrils. Curr Opin Plant Biol 11, 641-646. Macias, M.J., Musacchio, A., Ponstingl, H., Nilges, M., Saraste, M., and Oschkinat, H. (1994). Structure of the pleckstrin homology domain from β-spectrin. Nature 369, 675-677. Marshall, A.J., Krahn, A.K., Ma, K., Duronio, V., and Hou, S. (2002). TAPP1 and TAPP2 are targets of phosphatidylinositol 3-kinase signaling in B cells: sustained plasma membrane recruitment triggered by the B-cell antigen receptor. Mol Cell Biol 22, 5479-5491. Martinez-Rucobo, F.W., Eckhardt-Strelau, L., and Terwisscha van Scheltinga, A.C. (2009). Yeast chitin synthase 2 activity is modulated by proteolysis and phosphorylation. Biochem J 417, 547-554. Matthysse, A.G., Holmes, K.V., and Gurlitz, R.H. (1981). Elaboration of cellulose fibrils by Agrobacterium tumefaciens during attachment to carrot cells. J Bacteriol 145, 583- 595. Matthysse, A.G., White, S., and Lightfoot, R. (1995a). Genes required for cellulose synthesis in Agrobacterium tumefaciens. J Bacteriol 177, 1069-1075. Matthysse, A.G., Thomas, D.L., and White, A.R. (1995b). Mechanism of cellulose synthesis in Agrobacterium tumefaciens. J Bacteriol 177, 1076-1081. Matthysse, A.G., Deora, R., Mishra, M., and Torres, A.G. (2008). Polysaccharides cellulose, poly-β-1,6-n-acetyl-D-glucosamine, and colanic acid are required for optimal binding of Escherichia coli O157:H7 strains to alfalfa sprouts and K-12 strains to plastic but not for binding to epithelial cells. Appl Environ Microbiol 74, 2384-2390. Matthysse, A.G., Deschet, K., Williams, M., Marry, M., White, A.R., and Smith, W.C. (2004). A functional cellulose synthase from Ascidian epidermis. Proc Natl Acad Sci U S A 101, 986-991. Mayer, B.J., Ren, R.B., Clark, K.L., and Baltimore, D. (1993). A putative modular domain present in diverse signaling proteins. Cell 73, 629-630.

Mayer, R., Ross, P., Weinhouse, H., Amikam, D., Volman, G., Ohana, P., Calhoon, R.D., Wong, H.C., Emerick, A.W., and Benziman, M. (1991). Polypeptide composition of bacterial cyclic diguanylic acid-dependent cellulose synthase and the occurrence of immunologically cross-reacting proteins in higher plants. Proc Natl Acad Sci U S A 88, 5472-5476. Mazur, P., Morin, N., Baginsky, W., el-Sherbeini, M., Clemas, J.A., Nielsen, J.B., and Foor, F. (1995). Differential expression and function of two homologous subunits of yeast 1,3-β-D-glucan synthase. Mol Cell Biol 15, 5671-5681. Meijer, H.J., and Govers, F. (2006). Genomewide analysis of phospholipid signaling genes in Phytophthora spp.: novelties and a missing link. Mol Plant Microbe Interact 19, 1337-1347. Melida, H., Encina, A., Alvarez, J., Acebes, J.L., and Caparros-Ruiz, D. (2010). Unraveling the biochemical and molecular networks involved in maize cell habituation to the cellulose biosynthesis inhibitor dichlobenil. Mol Plant 3, 842-853. Melida, H., Garcia-Angulo, P., Alonso-Simon, A., Encina, A., Alvarez, J., and Acebes, J.L. (2009). Novel type II cell wall architecture in dichlobenil-habituated maize calluses. Planta 229, 617-631. Meyer, F.P. (1991). Aquaculture disease and health management. J Anim Sci 69, 4201-4208. Meyer, Y., and Hert, W. (1978). Chemical inhibition of cell wall formation and cytokinesis, but not of nuclear division, in protoplasts of Nicotiana tabacum L. cultivated in vitro. Planta 142, 253-262. Mitchell, H.J., and Hardham, A.R. (1999). Characterisation of the water expulsion vacuole in Phytophthora nicotianae zoospores. Protoplasma 206, 118-130. Montezinos, D., and Delmer, D.P. (1980). Characterization of inhibitors of cellulose synthesis in cotton fibers. Planta 148, 305-311. Morris, P.F., Schlosser, L.R., Onasch, K.D., Wittenschlaeger, T., Austin, R., and Provart, N. (2009). Multiple horizontal gene transfer events and domain fusions have created novel regulatory and metabolic networks in the Oomycete genome. PLoS One 4, e6133. Mort-Bontemps, M., Gay, L., and Fevre, M. (1997). CHS2, a chitin synthase gene from the Oomycete Saprolegnia monoica. Microbiol 143, 2009-2020. Mutwil, M., Debolt, S., and Persson, S. (2008). Cellulose synthesis: a complex complex. Curr Opin Plant Biol 11, 252-257. Nakagawa, N., and Sakurai, N. (1998). Increase in the amount of celA1 protein in tobacco BY-2 cells by a cellulose biosynthesis inhibitor, 2,6-dichlorobenzonitrile. Plant Cell Physiol 39, 779-785. Nakashima, K., Yamada, L., Satou, Y., Azuma, J., and Satoh, N. (2004). The evolutionary origin of animal cellulose synthase. Dev Genes Evol 214, 81-88. Neish, G.A. (1977). Observations of Saprolegniosis on adult sockeye salmon, Oncorhynchus nerka (Walbaum). J Fish Biol 10, 513-&. Nelson, N. (1944). A photometric adaptation of the somogyi method for the determination of glucose. J. Biol. Chem. 153, 375-380. Nicol, F., His, I., Jauneau, A., Vernhettes, S., Canut, H., and Hofte, H. (1998). A plasma membrane-bound putative endo-1,4-β-D-glucanase is required for normal wall assembly and cell elongation in Arabidopsis. EMBO J 17, 5563-5576. Niederkorn, J.Y., Alizadeh, H., Leher, H., and McCulley, J.P. (1999). The pathogenesis of Acanthamoeba keratitis. Microbes Infect 1, 437-443. Nobles, D.R., Romanovicz, D.K., and Brown, R.M. (2001). Cellulose in cyanobacteria. Origin of vascular plant cellulose synthase? J Plant Physiol 127, 529-542.

Nodet, P., Capellano, A., and Fevre, M. (1990 a). Morphogenetic effects of congo red on hyphal growth and cell wall development of the fungus Saprolegnia monoica. J Gen Microbiol 136, 303-310. Nodet, P., Girard, V., and Fevre, M. (1990 b). Congo red inhibits in vitro β-glucan synthases of Saprolegnia. FEMS 69, 225-228. Nühse, T.S., Stensballe, A., Jensen, O.N., and Peck, S.C. (2004). Phosphoproteomics of the Arabidopsis plasma membrane and a new phosphorylation site database. Plant Cell 16, 2394-2405. Pagant, S., Bichet, A., Sugimoto, K., Lerouxel, O., Desprez, T., McCann, M., Lerouge, P., Vernhettes, S., and Höfte, H. (2002). KOBITO1 encodes a novel plasma membrane protein necessary for normal synthesis of cellulose during cell expansion in Arabidopsis. Plant Cell 14, 2001-2013. Paquin, B., Laforest, M.J., Forget, L., Roewer, I., Wang, Z., Longcore, J., and Lang, B.F. (1997). The fungal mitochondrial genome project: evolution of fungal mitochondrial genomes and their gene expression. Curr Genet 31, 380-395. Paredez, A.R., Somerville, C.R., and Ehrhardt, D.W. (2006). Visualization of cellulose synthase demonstrates functional association with microtubules. Science 312, 1491- 1495. Paredez, A.R., Persson, S., Ehrhardt, D.W., and Somerville, C.R. (2008). Genetic evidence that cellulose synthase activity influences microtubule cortical array organization. Plant Physiol 147, 1723-1734. Paterson, H.F., Savopoulos, J.W., Perisic, O., Cheung, R., Ellis, M.V., Williams, R.L., and Katan, M. (1995). Phospholipase C δ 1 requires a pleckstrin homology domain for interaction with the plasma membrane. Biochem J 312 661-666. Pear, J.R., Kawagoe, Y., Schreckengost, W.E., Delmer, D.P., and Stalker, D.M. (1996). Higher plants contain homologs of the bacterial celA genes encoding the catalytic subunit of cellulose synthase. Proc Natl Acad Sci U S A 93, 12637-12642. Peng, L.C., Kawagoe, Y., Hogan, P., and Delmer, D. (2002). Sitosterol-β-glucoside as primer for cellulose synthesis in plants. Science 295, 147-150. Pérez, S., and Mazeau, K. (2005). Conformations, structures and morphologies of cellulose. In Polysaccharides: Structural diversity and functional versatility, D. S, ed (Montecillo, NY), pp. 41-69. Persson, S., Paredez, A., Carroll, A., Palsdottir, H., Doblin, M., Poindexter, P., Khitrov, N., Auer, M., and Somerville, C.R. (2007). Genetic evidence for three unique components in primary cell-wall cellulose synthase complexes in Arabidopsis. Proc Natl Acad Sci U S A 104, 15566-15571. Phillips, A.J., Anderson, V.L., Robertson, E.J., Secombes, C.J., and van West, P. (2008). New insights into animal pathogenic Oomycetes. Trends Microbiol 16, 13-19. Qadota, H., Python, C.P., Inoue, S.B., Arisawa, M., Anraku, Y., Zheng, Y., Watanabe, T., Levin, D.E., and Ohya, Y. (1996). Identification of yeast Rho1p GTPase as a regulatory subunit of 1,3-β-glucan synthase. Science 272, 279-281. Rajangam, A.S., Kumar, M., Aspeborg, H., Guerriero, G., Arvestad, L., Pansri, P., Brown, C.J., Hober, S., Blomqvist, K., Divne, C., Ezcurra, I., Mellerowicz, E., Sundberg, B., Bulone, V., and Teeri, T.T. (2008). MAP20, a microtubule-associated protein in the secondary cell walls of hybrid aspen, is a target of the cellulose synthesis inhibitor 2,6-dichlorobenzonitrile. Plant Physiol 148, 1283-1294. Rameh, L.E., Arvidsson, A., Carraway, K.L., 3rd, Couvillon, A.D., Rathbun, G., Crompton, A., VanRenterghem, B., Czech, M.P., Ravichandran, K.S., Burakoff, S.J., Wang, D.S., Chen, C.S., and Cantley, L.C. (1997). A comparative analysis of

the phosphoinositide binding specificity of pleckstrin homology domains. J Biol Chem 272, 22059-22066. Rinaudo, M. (2006). Chitin and chitosan: Properties and applications. Progr Poly Sci 31, 603-632. Rizzo, D.M., Garbelotto, M., Davidson, J.M., Slaughter, G.W., and Koike, S.T. (2002). Phytophthora ramorum as the cause of extensive mortality of Quercus spp. and Lithocarpus densiflorus in California. Plant Dis 86, 205-214. Roncero, C. (2002). The genetic complexity of chitin synthesis in fungi. Current Genetics 41, 367-378. Ronquist, F., and Huelsenbeck, J.P. (2003). MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19, 1572-1574. Ross, P., Mayer, R., and Benziman, M. (1991). Cellulose biosynthesis and function in bacteria. Microbiol Rev 55, 35-58. Rudsander, U., Denman, S., Raza, S., and Teeri, T.T. (2003). Molecular features of family GH9 cellulases in hybrid Aspen and the filamentous fungus Phanerochaete chrysosporium. J Appl Glycosci 50, 253-256. Ruiz-Herrera, J., and Ortiz-Castellanos, L. (2009). Analysis of the phylogenetic relationships and evolution of the cell walls from yeasts and fungi. FEMS Yeast Res 10, 225-243. Ruiz-Herrera, J., Gonzalez-Prieto, J.M., and Ruiz-Medrano, R. (2002). Evolution and phylogenetic relationships of chitin synthases from yeasts and fungi. FEMS Yeast Res 1, 247-256. Römling, U. (2002). Molecular biology of cellulose production in bacteria. Res Microbiol 153, 205-212. Sabba, R.P., and Vaughn, K.C. (1999). Herbicides that inhibit cellulose biosynthesis. Weed Sci 47, 757–763. Saxena, I.M., and Brown, R.M. (1997). Identification of cellulose synthase(s) in higher plants: Sequence analysis of processive β-glycosyltransferases with the common motif 'D, D, D35Q(R,Q)XRW'. Cellulose 4, 33-49. Saxena, I.M., and Brown, R.M., Jr. (2000). Cellulose synthases and related enzymes. Curr Opin Plant Biol 3, 523-531. Saxena, I.M., Lin, F.C., and Brown, R.M., Jr. (1990). Cloning and sequencing of the cellulose synthase catalytic subunit gene of Acetobacter xylinum. Plant Mol Biol 15, 673-683. Saxena, I.M., Brown, R.M., Jr., and Dandekar, T. (2001). Structure-function characterization of cellulose synthase: relationship to other glycosyltransferases. Phytochemistry 57, 1135-1148. Saxena, I.M., Kudlicka, K., Okuda, K., and Brown, R.M. (1994). Characterization of genes in the cellulose synthesizing operon (Acs operon) of Acetobacter xylinum - Implications for cellulose crystallization. J Bacteriol 176, 5735-5752. Saxena, I.M., Brown, R.M., Fèvre, M., Geremia, R.A., and Henrissat, B. (1995). Multidomain architecture of β-glycosyl transferases - Implications for mechanism of action. J Bacteriol 177, 1419-1424. Sburlati, A., and Cabib, E. (1986). Chitin synthetase 2, a presumptive participant in septum formation in Saccharomyces cerevisiae. J Biol Chem 261, 15147-15152. Scheible, W.R., Eshed, R., Richmond, T., Delmer, D., and Somerville, C. (2001). Modifications of cellulose synthase confer resistance to isoxaben and thiazolidinone herbicides in Arabidopsis Ixr1 mutants. Proc Natl Acad Sci U S A 98, 10079-10084. Schindelman, G., Morikami, A., Jung, J., Baskin, T.I., Carpita, N.C., Derbyshire, P., McCann, M.C., and Benfey, P.N. (2001). COBRA encodes a putative GPI-anchored

protein, which is polarly localized and necessary for oriented cell expansion in Arabidopsis. Gene Dev 15, 1115-1127. Schrick, K. (2004). A link between sterol biosynthesis, the cell-wall, and cellulose in Arabidopsis. Plant J 38, 562-562. Schumann, G.L. (1991). The Irish potato famine and the birth of plant pathology In Plant diseases: their biology and social impact (St. Paul, Minnesota: APS Press), pp. 1-20. Shahinian, S., and Bussey, H. (2000). β-1,6-glucan synthesis in Saccharomyces cerevisiae. Mol Microbiol 35, 477-489. Shaw, J.A., Mol, P.C., Bowers, B., Silverman, S.J., Valdivieso, M.H., Duran, A., and Cabib, E. (1991). The function of chitin synthase-2 and synthase-3 in the Saccharomyces cerevisiae cell cycle. J of Cell Biology 114, 111-123. Shedletzky, E., Shmuel, M., Delmer, D.P., and Lamport, D.T. (1990). Adaptation and growth of tomato cells on the herbicide 2,6-dichlorobenzonitrile leads to production of unique cell walls virtually lacking a cellulose-xyloglucan network. Plant Physiol 94, 980-987. Shedletzky, E., Shmuel, M., Trainin, T., Kalman, S., and Delmer, D. (1992). Cell wall structure in cells adapted to growth on the cellulose-synthesis inhibitor 2,6- dichlorobenzonitrile : a comparison between two dicotyledonous plants and a graminaceous monocot. Plant Physiol 100, 120-130. Si-Ammour, A., Mauch-Mani, B., and Mauch, F. (2003). Quantification of induced resistance against Phytophthora species expressing GFP as a vital marker: β- aminobutyric acid but not BTH protects potato and Arabidopsis from infection. Mol Plant Pathol 4, 237-248. Sinnott, M.L. (1990). Catalytic mechanisms of enzymatic glycosyl transfer. Chem Rev 90, 1171-1202. Song, D., Shen, J., and Li, L. (2010). Characterization of cellulose synthase complexes in Populus xylem differentiation. New Phytol 187, 777-790. Southern, E.M. (1975). Detection of specific sequences among DNA fragments separated by gel electrophoresis. J Mol Biol 98, 503-517. Standal, R., Iversen, T.G., Coucheron, D.H., Fjaervik, E., Blatny, J.M., and Valla, S. (1994). A new gene required for cellulose production and a gene encoding cellulolytic activity in Acetobacter xylinum are colocalized with the bcs operon. J Bacteriol 176, 665-672. Stasinopoulos, S.J., Fisher, P.R., Stone, B.A., and Stanisich, V.A. (1999). Detection of two loci involved in (1->3)-β-glucan (curdlan) biosynthesis by Agrobacterium sp. ATCC31749, and comparative sequence analysis of the putative curdlan synthase gene. Glycobiology 9, 31-41. Stone, B.A., and Clarke, A.E. (1992). Chemistry and biology of (1−>3)-β-glucans (Victoria, Australia: La Trobe University Press), pp. 431-491. Strynadka, N.C.J., Lovering, A.L., Lin, L.Y.C., Sewell, E.W., Spreter, T., and Brown, E.D. (2010). Structure of the bacterial teichoic acid polymerase TagF provides insights into membrane association and catalysis. Nat Struct Mol Biol 17, 582-U581. Suzuki, S., Li, L., Sun, Y.H., and Chiang, V.L. (2006). The cellulose synthase gene superfamily and biochemical functions of xylem-specific cellulose synthase like genes in Populus trichocarpa. Plant Physiol 142, 1233-1245. Swissa, M., Aloni, Y., Weinhouse, H., and Benizman, M. (1980). Intermediary steps in Acetobacter xylinum cellulose synthesis - studies with whole cells and cell-free preparations of the wild-type and a cellulose-less mutant. J Bacteriol 143, 1142-1150.

Takeshita, N., Ohta, A., and Horiuchi, H. (2005). CsmA, a class V chitin synthase with a myosin motor-like domain, is localized through direct interaction with the actin cytoskeleton in Aspergillus nidulans. Mol Biol Cell 16, 1961-1970. Taylor, N.G., Laurie, S., and Turner, S.R. (2000). Multiple cellulose synthase catalytic subunits are required for cellulose synthesis in Arabidopsis. Plant Cell 12, 2529-2540. Taylor, N.G., Howells, R.M., Huttly, A.K., Vickers, K., and Turner, S.R. (2003). Interactions among three distinct CesA proteins essential for cellulose synthesis. Proc Natl Acad Sci U S A 100, 1450-1455. Timmers, J., Vernhettes, S., Desprez, T., Vincken, J.P., Visser, R.G.F., and Trindade, L.M. (2009). Interactions between membrane-bound cellulose synthases involved in the synthesis of the secondary cell wall. FEBS Lett 583, 978-982. Torto-Alalibo, T., Tian, M., Gajendran, K., Waugh, M.E., van West, P., and Kamoun, S. (2005). Expressed sequence tags from the Oomycete fish pathogen Saprolegnia parasitica reveal putative virulence factors. BMC Microbiol 5, 46. Touhara, K., Inglese, J., Pitcher, J.A., Shaw, G., and Lefkowitz, R.J. (1994). Binding of G protein βγ-subunits to pleckstrin homology domains. J Biol Chem 269, 10217-10220. Tyers, M., Rachubinski, R.A., Stewart, M.I., Varrichio, A.M., Shorr, R.G., Haslam, R.J., and Harley, C.B. (1988). Molecular cloning and expression of the major protein kinase C substrate of platelets. Nature 333, 470-473. Tyler, B.M. (2002). Molecular basis of recognition between Phytophthora pathogens and their hosts. Annu Rev Phytopathol 40, 137-167. Tyler, B.M., Tripathy, S., Zhang, X., Dehal, P., Jiang, R.H., Aerts, A., Arredondo, F.D., Baxter, L., Bensasson, D., and al., e. (2006a). Phytophthora genome sequences uncover evolutionary origins and mechanisms of pathogenesis. Science 313, 1261- 1266. Tyler, B.M., Tripathy, S., Zhang, X.M., Dehal, P., Jiang, R.H.Y., Aerts, A., Arredondo, F.D., Baxter, L., Bensasson, D., Beynon, J.L., Chapman, J., Damasceno, C.M.B., Dorrance, A.E., Dou, D.L., Dickerman, A.W., Dubchak, I.L., Garbelotto, M., Gijzen, M., Gordon, S.G., Govers, F., Grunwald, N.J., Huang, W., Ivors, K.L., Jones, R.W., Kamoun, S., Krampis, K., Lamour, K.H., Lee, M.K., McDonald, W.H., Medina, M., Meijer, H.J.G., Nordberg, E.K., Maclean, D.J., Ospina- Giraldo, M.D., Morris, P.F., Phuntumart, V., Putnam, N.H., Rash, S., Rose, J.K.C., Sakihama, Y., Salamov, A.A., Savidor, A., Scheuring, C.F., Smith, B.M., Sobral, B.W.S., Terry, A., Torto-Alalibo, T.A., Win, J., Xu, Z.Y., Zhang, H.B., Grigoriev, I.V., Rokhsar, D.S., and Boore, J.L. (2006b). Phytophthora genome sequences uncover evolutionary origins and mechanisms of pathogenesis. Science 313, 1261-1266. Uchida, Y., Shimmi, O., Sudoh, M., Arisawa, M., and Yamada-Okabe, H. (1996). Characterization of chitin synthase 2 of Saccharomyces cerevisiae. II: Both full size and processed enzymes are active for chitin synthesis. J Biochem 119, 659-666. Ude, S., Arnold, D.L., Moon, C.D., Timms-Wilson, T., and Spiers, A.J. (2006). Biofilm formation and cellulose expression among diverse environmental Pseudomonas isolates. Environ Microbiol 8, 1997-2011. Valdivieso, M.H., Mol, P.C., Shaw, J.A., Cabib, E., and Duran, A. (1991). Cal1, a gene required for activity of chitin synthase-3 in Saccharomyces cerevisiae. J Cell Biol 114, 101-109. Walker, C.A., and van West, P. (2007). Zoospore development in the Oomycetes. Fung Biol Rev 21, 10-18. van der Vaart, J.M., te Biesebeke, R., Chapman, J.W., Klis, F.M., and Verrips, C.T. (1996). The β-1, 6-glucan containing side-chain of cell wall proteins of

Saccharomyces cerevisiae is bound to the glycan core of the GPI moiety. FEMS Lett 145, 401-407. van West, P. (2006). Saprolegnia parasitica, an Oomycete pathogen with a fishy appetite: new challenges for an old problem. Mycologist 20, 99-104. Werner, S., Steiner, U., Becher, R., Kortekamp, A., Zyprian, E., and Deising, H.B. (2002). Chitin synthesis during in planta growth and asexual propagation of the cellulosic Oomycete and obligate biotrophic grapevine pathogen Plasmopara viticola. FEMS Microbiol Lett. 208, 169-173. Whisson, S.C., Avrova, A.O., Van West, P., and Jones, J.T. (2005). A method for double- stranded RNA-mediated transient gene silencing in Phytophthora infestans. Mol Plant Pathol 6, 153-163. Willoughby, L.G., and Pickering, A.D. (1977). Viable Saprolegniaceae spores on epidermis of salmonid fish Salmo trutta and Salvelinus alpinus. Trans Brit Mycol Soc 68, 91-95. Wong, H.C., Fear, A.L., Calhoon, R.D., Eichinger, G.H., Mayer, R., Amikam, D., Benziman, M., Gelfand, D.H., Meade, J.H., Emerick, A.W., and et al. (1990). Genetic organization of the cellulose synthase operon in Acetobacter xylinum. Proc Natl Acad Sci U S A 87, 8130-8134. Wood, P.J. (1980). Specificity in the interaction of direct dyes with polysaccharides. Carbohydr Res 85, 271-287. Xu, S.L., Rahman, A., Baskin, T.I., and Kieber, J.J. (2008). Two leucine-rich repeat receptor kinases mediate signaling, linking cell wall biosynthesis and ACC synthase in Arabidopsis. Plant Cell 20, 3065-3079. Yao, L., Kawakami, Y., and Kawakami, T. (1994). The pleckstrin homology domain of Bruton tyrosine kinase interacts with protein kinase C. Proc Natl Acad Sci U S A 91, 9175-9179. Yao, L., Janmey, P., Frigeri, L.G., Han, W., Fujita, J., Kawakami, Y., Apgar, J.R., and Kawakami, T. (1999). Pleckstrin homology domains interact with filamentous actin. J Biol Chem 274, 19752-19761. Yoon, H.S., Hajduk, P.J., Petros, A.M., Olejniczak, E.T., Meadows, R.P., and Fesik, S.W. (1994). Solution structure of a pleckstrin homology domain. Nature 369, 672- 675. Zheng, J., Chen, R.H., Corblan-Garcia, S., Cahill, S.M., Bar-Sagi, D., and Cowburn, D. (1997). The solution structure of the pleckstrin homology domain of human SOS1. A possible structural role for the sequential association of diffuse B cell lymphoma and pleckstrin homology domains. J Biol Chem 272, 30340-30344.