Characterization of Specific Domains of the Cellulose and Chitin Synthases from Pathogenic Oomycetes

Characterization of specific domains of the cellulose and chitin synthases from pathogenic oomycetes

Christian Brown

Doctoral thesis in Biotechnology

School of Biotechnology Royal Institute of Technology Stockholm, Sweden 2015

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

Printed by Universitetsservice US-AB Drottning Kristinas väg 53B SE-100 44 Stockholm Sweden

TRITA-BIO Report 2015:15 ISBN 978-91-7595-690-9 ISSN 1654-2312

Cover: NMR structure of Saprolegnia monoica MIT domain from chitin synthase 1. Christian Brown (2015) Characterization of specific domains of the cellulose and chitin synthases from pathogenic oomycetes. Doctoral Thesis in Biotechnology. Royal Institute of Technology (KTH), Division of Glycoscience, Stockholm, Sweden. TRITA-BIO Report 2015:15 ISBN 978-91-7595-690-9 ISSN 1654-2312

ABSTRACT

Some oomycetes species are severe pathogens of fish or crops. As such, they are responsible for important losses in the aquaculture industry as well as in agriculture. Saprolegnia parasitica is a major concern in aquaculture as there is currently no method available for controlling the diseases caused by this microorganism. The cell wall is an extracellular matrix composed essentially of polysaccharides, whose integrity is required for oomycete viability. Thus, the enzymes involved in the biosynthesis of cell wall components, such as cellulose and chitin synthases, represent ideal targets for disease control. However, the biochemical properties of these enzymes are poorly understood, which limits our capacity to develop specific inhibitors that can be used for blocking the growth of pathogenic oomycetes.

In our work, we have used Saprolegnia monoica as a model species for oomycetes to characterize two types of domains that occur specifically in oomycete carbohydrate synthases: the Pleckstrin Homology (PH) domain of a cellulose synthase and the so-called ‘Microtubule Interacting and Trafficking’ (MIT) domain of chitin synthases. In addition, the chitin synthase activity of the oomycete phytopathogen Aphanomyces euteiches was characterized in vitro using biochemical approaches.

The results from our in vitro investigations revealed that the PH domain of the oomycete cellulose synthase binds to phosphoinositides, microtubules and F-actin. In addition, cell biology approaches were used to demonstrate that the PH domain co-localize with F-actin in vivo. The structure of the MIT domain of chitin synthase (CHS) 1 was solved by NMR. In vitro binding assays performed on recombinant MIT domains from CHS 1 and CHS 2 demonstrated that both proteins strongly interact with phosphatidic acid in vitro. These results were further supported by in silico data where biomimetic membranes composed of different phospholipids were designed for interaction studies. The use of a yeast-two-hybrid approach suggested that the MIT domain of CHS 2 interacts with the delta subunit of Adaptor Protein 3, which is involved in protein trafficking. These data support a role of the MIT domains in the cellular targeting of CHS proteins. Our biochemical data on the characterization of the chitin synthase activity of A. euteiches suggest the existence of two distinct enzymes responsible for the formation of water soluble and insoluble chitosaccharides, which is consistent with the existence of two putative CHS genes in the genome of this species.

Altogether our data support a role of the PH domain of cellulose synthase and MIT domains of CHS in membrane trafficking and cellular location.

Keywords: Cellulose biosynthesis; chitin biosynthesis; cellulose synthase genes; chitin synthase genes; oomycetes; Saprolegnia monoica; Microtubule Interacting and Trafficking (MIT) domain; Pleckstrin Homology (PH) domain

SAMMANFATTING

En del arter av oomyceter är allvarliga patogener för fisk eller grödor. De kan därför anses vara ansvariga för väsentliga förluster för både vatten- och lantbruk. Saprolegnia parasitica är ett stort problem inom vattenbruket, då det för närvarande inte finns någon metod för att hindra sjukdomarna som denna parasit orsakar. Cellväggen är en extracellulär matrix som till största del består av polysackarider, vars integritet krävs för att oomyceterna ska kunna överleva. Därför är enzymerna som är aktiva i biosyntesen av cellväggskomponenter, som till exempel cellulosa- och kitinsyntaser, perfekta måltavlor för smittskyddskontroll. De biokemiska egenskaperna hos dessa enzymer är dock dåligt klarlagda, vilket begränsar vår möjlighet till att utveckla specifika inhibitorer som kan användas för att hindra tillväxten av patogeniska oomyceter.

I vårt arbete har vi använt Saprolegnia monoica som en modell för oomyceter för att karaktärisera två typer av domäner som förekommer specifikt i syntas av kolhydrater: Pleckstrinhomologidomänen (PH- domänen) hos en cellulosasyntas, och den så kallade "Microtubule Interacting and Trafficking"- domänen, eller MIT-domänen, hos kitinsyntas. Dessutom karaktäriserades kitinsyntasen hos oomycetfytopatologen Aphanomyces euteiches in vitro med hjälp av biokemiska metoder.

Resultaten från vår in vitro-forskning visade att PH-domänen hos syntasen av oomycetcellulosan binder med fosfoinositider, mikrotubuli och F-aktin. Dessutom användes cellbiologiska metoder för att visa att PH-domänen samlokaliserar med F-aktin in vivo. Strukturen av MIT-domänen hos kitinsyntasen (CHS) 1 löstes med NMR. Bindingsanalyser in vitro som genomfördes på rekombinanta MIT-domäner från CHS 1 och CHS 2 visade att båda proteinerna reagerar starkt med fosfatidinsyra in vitro. Resultaten stärks även av in silico-data där biometriska membran som bestod av olika fosfolipider utformades specifikt för interaktionsstudier. Användningen av en jäst-två-hybridmetod (Y2H) visade att MIT-domänen hos CHS 2 reagerar med delta-subenheten hos Adaptorprotein 3, som är aktiv i proteinförflyttning. Dessa data stödjer MIT-domänernas roll i cellinriktningen av CHS- proteiner. Våra biokemiska data över karaktäriseringen av kitinsyntasen hos A. euteiches tyder på att det finns två distinkta enzymer som är ansvariga för bildandet av vattenlösliga och icke-vattenlösliga kitosackarider, vilket stämmer överens med förekomsten av två förmodade CHS-gener i artens arvsmassa.

Sammanfattningsvis stöder våra data att PH-domänen har en roll i cellulosasyntas, och MIT-domäner hos CHS har en roll i membranförflyttning och cellulär lokalisering

Nyckelord: Biosyntes av cellulosa; biosyntes av kitin; cellulose synthase genes; chitin synthase genes; oomyceter; Saprolegnia monoica; Microtubule Interacting and Trafficking domain (MIT); Pleckstrinhomologidomän (PH-domän)

LIST OF PUBLICATIONS

Publication I Brown, C., Leijon, F., and Bulone, V. (2012). Radiometric and spectrophotometric in vitro assays of glycosyltransferases involved in plant cell wall carbohydrate biosynthesis. Nat Protoc. 7, 1634-1650.

Publication II Fugelstad, J*., Brown, C*., Hukasova, E., Sundqvist, G., Lindqvist, A., and Bulone, V. (2012). Functional characterization of the pleckstrin homology domain of a cellulose synthase from the Oomycete Saprolegnia monoica. Biochem. Biophys. Res. Comm. 417, 1248-1253.

Publication III Brown, C*., Szpryngiel, S*., Kuang, G., Srivastava, V., Weihua, Ye., McKee, L.S., Tu, Y., Mäler, L., and Bulone, V. (2015). Structural and functional characterization of the “Microtubule Interacting and Trafficking” domains of two oomycetes chitin synthases. Submitted to Biochem. J.

Publication IV Kuang, G., Liang, L., Brown, C., Wang, Q., Bulone, V., and Tu, Y. (2015). Insight into the adsorption profiles of the MIT domain of a chitin synthase from Oomycete Saprolegnia monoica on the POPA and POPC membranes by molecular dynamics simulation studies. Submitted to Phys. Chem. Chem. Phys.

Publication V Bottin, A., Brown, C., Melida, H., and Bulone, V. (2015). The phytopathogenic oomycete Aphanomyces euteiches contains two distinct N acetylglucosaminyltransferase activities that form chitin-like saccharides in vitro. Manuscript.

*Both authors contributed equally to the work

iii

THE AUTHOR’S CONTRIBUTION

Publication I: Christian Brown performed complementary optimization experiments and contributed to the writing of the experimental section and the preparation of the figures.

Publication II: Christian Brown performed the phosphoinositide binding assays, the in vitro binding experiments to F-actin and microtubules, prepared figures and wrote the Materials and Methods sections related to these experiments.

Publication III: Christian Brown performed the recombinant expression, purification and biochemical characterization of the MIT domains of the oomycete chitin synthases and contributed to the writing of all sections of the manuscript.

Publication IV: Christian Brown contributed complementary experiments to the in silico approaches and assisted in the design of the models used for modeling. He prepared some of the figures and contributed to the writing of the manuscript.

Publication V: Christian Brown performed the biochemical work with co-author Dr Arnaud Bottin and contributed to the preparation of the figures and writing of the experimental section.

RELATED PUBLICATIONS NOT INCLUDED IN THE THESIS

Rajangam, A.S., Kumar, M., Aspeborg, H., Guerriero, G., Arvestad, L., Pansri, P., Brown, C., Hober, S., Blomqvist, K., Divne, C., Ezcurra, I., Sundberg, B., Mellerowicz, E., Bulone, V., and Teeri, T.T. (2008). PttMAP20, a novel microtubule- associated protein in the secondary cell wall forming tissues of Populus tremula x tremuloides. Plant Physiol. 148, 1283-1294

Butchosa, N., Brown, C., Larsson, T., Bulone, V., and Zhou, Q. (2013). Nanocomposites of bacterial cellulose nanofibers and chitin nanocrystals: fabrication, characterization and bactericidal activity. Green Chem. 15, 3404-3413

Table of contents

Abstract……………………………………………………………………………………...... I Sammanfattning……………………………………………………………………………………..II List of publications…………………………………………………………………………………III The author’s contribution…………………………………………………………………………IV Related publications not included in the thesis………………………………………………IV Table of contents……………………………………………………………………………………V List of figures……………………………………………………………………………………….VII List of tables………………………………………………………………………………………..VIII

1 Introduction ...... 1 1.1 Oomycetes and their importance ...... 1 1.2 Life cycle ...... 4 1.3 The cell wall ...... 6 1.3.1 Oomycete cell walls ...... 6 1.3.2 Fungal and yeast cell walls ...... 9 1.4 Carbohydrate-Active enZymes (CAZymes) ...... 13 1.5 Chitin ...... 16 1.5.1 Chitin structure ...... 16 1.6 Chitin biosynthesis ...... 18 1.6.1 Main process ...... 18 1.6.2 Chitin synthases ...... 21 1.6.3 Post-translational regulation of chitin synthases ...... 26 1.6.3.1 Phosphorylation ...... 26 1.6.3.2 Zymogenicity ...... 26 1.6.4 Targeting of chitin synthases to the plasma membrane ...... 27 1.6.4.1 Chitosomes ...... 27 1.6.4.2 Myosin motor-like domains of Chs ...... 28 1.6.4.3 MIT domains ...... 30 1.7 Cellulose ...... 30 1.7.1 Terminal complexes ...... 31 1.7.2 Cellulose structure and biosynthesis ...... 31 1.7.3 Cellulose synthase genes from oomycetes ...... 36 1.8 Overview of the secretory and endocytosis pathways of proteins ...... 39 v

1.8.1 Secretory pathway ...... 39 1.8.2 Clathrin-mediated endocytosis pathway ...... 42 1.9 Microtubule interacting and trafficking (MIT) domains ...... 43 1.10 MIT domains in carbohydrate-active enzymes and other types of proteins ...... 45 1.10.1 Carbohydrate-active enzymes ...... 45 1.10.2 Non-carbohydrate-active enzymes ...... 46 1.10.2.1 Spastin ...... 46 1.10.2.2 De-ubiquitinating enzymes...... 47 1.10.2.3 VPS4 (Vps4) ...... 47 1.11 Pleckstrin homology domains ...... 48 1.11.1 Background ...... 48 1.11.2 Structure of PH domains ...... 49 1.11.3 Phosphatidylinositol ...... 50 1.11.4 Function and mode of interaction of PH domains ...... 52 1.11.5 Classification of PH domains ...... 55 2 Aims of the present investigation ...... 56 2.1 The specific aims of the thesis were: ...... 56 3 Results and discussion ...... 57 3.1 Glycosyltransferase assays (Paper I) ...... 57 3.2 Characterization of the PH domain from SmCesA2 (Paper II) ...... 60 3.3 Biochemical and structural characterization of the SmChs1 MIT domain (Paper III) ....62 3.4 Modelling of the interactions between the MIT domain of SmChs1 (Sm_MIT1) and lipids (Paper IV) ...... 65 3.5 Characterization of two putative chitin synthases (AeChs1 and AeChs2) from microsomal fractions of Aphanomyces euteiches (Paper V) ...... 68 4 Conclusions and perspectives ...... 70 5 Acknowledgements ...... 75 6 References ...... 77

List of figures

Figure 1. Phylogenetic tree of eukaryotic organisms ...... 2 Figure 2. A male Chinook salmon infected by S. parasitica ...... 4 Figure 3. Life cycle of S. parasitica ...... 5 Figure 4. Most abundant types of glucans forming the oomycete cell wall...... 7 Figure 5. Cell wall model of C. albicans ...... 10 Figure 6. The GPI anchor structure ...... 13 Figure 7. Reaction catalysed by chitin synthases ...... 14 Figure 8. Proposed mechanism for inverting glycosyltransferases ...... 15 Figure 9. Proposed mechanism for retaining glycosyltransferases ...... 16 Figure 10. Representation of the different allomorphs of chitin ...... 17 Figure 11. Chitin structure and intramolecular hydrogen bonds ...... 17 Figure 12. The chitin synthesis pathway ...... 21 Figure 13. Domain comparison of oomycete Chs ...... 24 Figure 14. Hypothetical model describing cellulose biosynthesis ...... 33 Figure 15. Domain organisation of CesA from a diverse range of organisms ...... 38 Figure 16. Endocytosis process ...... 41 Figure 17. Representation of the adaptor protein complexes and their subunits ...... 42 Figure 18. Structure of the MIT domain from the human VPS4A protein ...... 44 Figure 19. Sequence alignment of MIT domains from oomycete Chs and other proteins ..... 46 Figure 20. Structure of the C-terminal PH domain of human pleckstrin ...... 49 Figure 21. Electrostatic potential of β-spectrin ...... 50 Figure 22. Structure of a phosphatidylinositol ...... 51 Figure 23. Specific distribution of phosphatidylinositides (PtdIns) in an animal cell ...... 52 Figure 24. Tools to vizualize and localize PtdIns in cell membranes ...... 54 Figure 25. Radiometric assays ...... 59 Figure 26. Lipid binding activity of SmCesA2PH ...... 61 Figure 27. Densitometric analysis of Sm_MIT1 and Sm_MIT2 binding on a Membrane ArrayTM...... 64 Figure 28. NMR structure of Sm_MIT1...... 65 Figure 29. Structure and sequence of Sm_MIT1 ...... 68 Figure 30. Hypothetical model representing possible targeting paths of Saprolegnia monoica chitin synthases to the plasma membrane (PM) and intracellular trafficking in hyphae ...... 72

vii List of tables

Table 1. Criteria characterizing the three major types of oomycete cell walls...... 8 Table 2. Classification of PH domains ...... 55 Table 3. Study of the different orientations (P1-P6) taken by Sm_MIT1 domain on the POPA membrane before and after adsorption...... 67

viii 1 Introduction ______

1.1 Oomycetes and their importance

As a consequence of the constant increase of the world population, the production of crops for human consumption has to be maximized. To sustain food needs, our use of the agricultural land and sea resources must be optimized and diseases brought by pathogens must be controlled. Pathogens and predators have been present since the beginning of life as they contribute to the ecological equilibrium of the planet. Oomycetes are a good example of a family of organisms that comprises plant and animal pathogens. They are eukaryotic microorganisms that can have a saprotroph and/or a necrotroph lifestyle. In the case of Saprolegnia species, they are considered as both saprotrophs and necrotrophs (Bruno and Wood, 1999; van West, 2006).

The classification of oomycetes has long been debated. They were previously classified in the kingdom Fungi because they share some similarities with fungal microorganisms, e.g. their mode of absorption of nutrients, reproduction by spores and filamentous growth. However, they also exhibit some clear differences from the fungi, in particular, the presence of septae is rare in oomycetes, which makes the hyphae typically multinucleate; the nuclei of vegetative cells are diploid and the cell wall composition is different from that of true fungi (Hardham, 2007). Phylogenetic studies and cell wall analyses have led to the conclusion that oomycetes are unrelated to true fungi (Baldauf et al., 2000). They are indeed more closely related to diatoms and brown algae than to fungi, and are currently classified in the Straminopila within the kingdom of Chromista (Cavalier-Smith and Chao 2006) (Figure 1).

Figure 1. Phylogenetic tree of eukaryotic organisms (adapted from Gijzen, 2009).

Some oomycete pathogens are well-known in agriculture and aquaculture industries where they cause significant economical losses (Meyer, 1991; Bruno and Wood, 1999). One of the well known plant pathogens, Phytophthora infestans, is the agent of potato late blight, which was responsible for the famine in Ireland in the 18th century. This species belongs to the Peronosporales order which comprises pathogens that infect not only plants, but also algae, protists, arthropods and vertebrates (Phillips et al., 2008). The oomycete Saprolegnia parasitica, a water mould, is an endemic fish pathogen found in fresh water all around the world, causing saprolegniosis to salmon, trout species and catfish (Figure 2). Control of this pathogen is important as it is causing high losses in the aquaculture industry and in natural habitats. For example, the salmon aquaculture business from different countries is losing many millions per year due to saprolegniosis only (Hussein & Hatai, 2002; van West, 2006). S. parasitica is also responsible for an estimated loss of €31 million each year in catfish farms in the USA (Bruno and Wood, 1999).

Until 2002, S. parasitica was controlled by using malachite green, where its major metabolite, Leucomalchite green accumulates in rainbow trout and catfish tissues

2 (Allen et al., 1994; Roybal et al., 1995; Plakas et al., 1996). This chemical is now banned worldwide due to its carcinogenic and toxicological effects. Aquaculture industries are currently facing a re-emergence of saprolegniosis. Moreover, S. parasitica is not only a problem for the fish farms, but it also infects wild salmon and therefore has a negative impact on fish populations in natural environments (Neitzel et al., 2004). Currently, very few chemicals are available to treat the eggs of salmonids and to make matter worse, no chemical can provide a sufficient protection after the eggs hatches (Fornerisa et al., 2003). A couple of ways to decrease the disease in aquaculture are to improve water quality by increasing the frequency of water being changed and also by adding NaCl (0,03M) as S. parasitica is inhibited at low concentrations of salt (Ali, 2005). Treatment of water with ozone can also prevent infection, but it does not cure infected fish. Formaldehyde and formalin formulations (a solution of formaldehyde + methanol + water) are used as a mean to reduce infections due to S. parasitica in fish farms (Gieseker et al., 2006), although formaldehyde is a toxic chemical that was reclassified in 2006 by the World Health Organisation (WHO) and International Agency for Research on Cancer (IARC) as “carcinogenic to humans” (WHO IARC, 2006) (Picón-Camacho et al., 2012). Therefore, new measures that have a low environmental impact and that are safe for humans are needed.

Bronopol (2-bromo-2-nitropropane-1,3-diol) is a broad-spectrum biocide that has been tested in the aquaculture industry against S. parasitica to replace malachite green for the treatment of salmonid fish and eggs (Pottinger and Day 1999; Branson, 2002). Before its application in aquaculture, bronopol was already used as a disinfectant for food production and water (Kumanova et al., 1989). Although bronopol has not been shown to have any negative effect on humans and has also been tested for environmental impact, its use in the UK needs an extra consent from the national environmental authorities (Picón-Camacho et al., 2012). Recent studies by Picón-Camacho et al. (2012) on rainbow trouts infected by Ichthyophthirius multifiliis (Ciliophora) showed that long exposures to low concentrations of bronopol decrease infection without affecting the trout’s health. Although the use of bronopol looks promising, it is costly and not easily available in all countries due to specific national regulations (Oono, 2007). Recent work on developing a vaccine against S.

3 parasitica to protect brown trout also seems promising and various strategies should be researched to control S. parasitica (Fregeneda-Grandes et al., 2007).

Another possible avenue is to target the cell wall of oomycetes, where the lack of proper cell wall formation in oomycetes could potentially inhibit their growth. Cell wall formation involves the activity of different types of enzymes, such as chitin and cellulose synthases that have been respectively shown to be important for the growth of Saprolegnia monoica (Guerriero et al., 2010; Fugelstad et al., 2009) and the infection process of P. infestans (Grenville-Briggs et al., 2008). Therefore, a better understanding of the cell wall biosynthesis and the properties of these enzymes can potentially provide new strategies to control pathogens as the cell walls are vital to cell survival (Pfaller et al., 2008). S. monoica, a species closely related to S. parasitica, has been used as a model in our laboratory to study cell wall formation. The composition of the cell walls of oomycetes is described in section 1.3.1.

Figure 2. A male Chinook salmon infected by S. parasitica (from van West, 2006). Infected fish are characterized by the presence of white and gray patches on their bodies (Lartseva, 1986).

1.2 Life cycle

The study of the life cycle of a pathogen is important to better understand its biology and in Figure 3, the life cycle of S. parasitica is illustrated as a representative example of oomycetes (after van West, 2006). The diploid life cycle of oomycetes can be divided in two phases i.e., sexual and asexual. The asexual phase allows the propagation and infection of the host. In S. parasitica, asexual cycle begins by

4 formation of sporangia on tips of the vegetative hyphae when the nutrients are depleted. The sporangia burst and release primary zoospores that quickly encyst and form primary cysts from which the secondary zoospores are released. These secondary zoospores can swim in water for longer periods of time and when they encyst, they form secondary cysts with special structures called “boat hooks” that can be described as filaments with hooks on their extremity. It has been proposed that the presence of these hairs enhances the attachment of the cysts to their host (Beakes, 1982; Pickering and Willoughby, 1982). The secondary cysts that do not find any host have the ability to release new zoospore; this is called polyplanetism (Figure 3). Once the secondary cyst is attached to the host, it will start the infection process by germination.

Figure 3. Life cycle of S. parasitica (adapted from van West, 2006).

5 1.3 The cell wall

1.3.1 Oomycete cell walls

The cell wall is typically described as a barrier that delimits the cell and determines its morphology. Cell morphology is also dependent on the turgor pressure exerted by the cytoplasm. The cell wall is involved in the control of exchanges between the cell and the environment. Carbohydrates are the main components of the cell wall and as such, largely contribute to the cell wall’s structure and properties. Glucans typically account for 50% to 90% of the dry weight of the cell wall (Bertke and Aronson, 1980). Carbohydrate-active enzymes are involved in the biosynthesis and modification of the cell wall during the life of a microorganism to respond to its needs (growth, reproduction, survival). Many enzymes are responsible for these processes, including glycosyltransferases (GTs), glycoside hydrolases and transglycosylases. In addition to carbohydrate-active enzymes, protein kinases and phosphatases may be involved in the morphogenesis of the hyphae. Only GTs are discussed in the context of this thesis (see section 1.4). The products formed by GTs are vital since oomycetes and fungi cannot survive without a cell wall. This can be exploited for disease control by identifying substances that block GTs and thus interfere with cell wall biosynthesis.

Early studies based on oomycete cell wall analyses showed that non-cellulosic glucans i.e. β-(1,3) and β-(1,6) glucans, as well as cellulose are the major carbohydrates of oomycete cell walls (Bartnicki-Garcia, 1968). β-(1,3)-Glucans are also found in plants in the form of callose but, as opposed to their oomycete counterparts (Figure 4A), plant β-(1,3)-glucans are not branched (Stone, 2006). Another type of β-glucan found in oomycete cell walls is β-(1,3;1,6) glucan (Mélida et al., 2013). It consists of a backbone of glucose residues linked by β-(1,3) glycosidic bonds. The backbone is also typically substituted at C-6 by chains that contain β- (1,3) and/or β-(1,6) linkages. As for the vast majority of natural polysaccharides, β- (1,3;1,6) glucans exhibit polydispersity in terms of molecular size, pattern and degree of branching.

6 As mentioned, cellulose is part of the oomycete cell wall and it is also a major component of plant cell walls, but, this polysaccharide is not found in fungi (Novaes- Ledieu, 1967) (Figure 4B). Cellulose is a water-insoluble homopolymer of glucose residues linked by β-(1,4) linkages (Figure 4B). The cellulose content varies between oomycete species. For example, the wall of Apodachlya (Saprolegniales) contains 4% cellulose (Sietsma, 1969), whereas the wall of Pythium (Peronosporales) contains up to 20% cellulose (Novaes-Ledieu et al., 1967). Cellulose in oomycetes is considered to have a similar function as chitin in fungi. It occurs in S. monoica hyphae as a network of non-oriented microfibrils and has been characterized as a poorly crystalline form of cellulose (type IV) by X-ray diffraction analysis (Bulone et al., 1992).

Figure 4. Most abundant types of glucans forming the oomycete cell wall. A. Structure of a β-(1,3) glucan with a β-(1,6) branch. B. Structure of a cellulose chain. The homopolymer consists of single units of glucose rotated 180º relative to each other to form β-(1,4) linkages. Cellobiose is sometimes considered as the repeating unit.

Another important cell wall carbohydrate is chitin, which consists of β-(1,4)-linked N- acetylglucosaminyl (GlcNAc) residues. This polysaccharide is typical of fungal cell walls but it has also been shown to occur in minute amounts in some oomycete

7 genera such as Apodachlya (Lin and Aronson, 1970), Leptomitus (Aronson and Lin, 1978), Apodachlyella (Bertke and Aronson, 1980) and Achlya (Campos-Takaki et al., 1982). In 1992, Bulone et al. fully characterized the chitin from cell walls of S. monoica. X-Ray diffraction analysis demonstrated the occurrence of a crystalline form of the α-type, while, electron microscopy revealed a granular morphology in hyphae. The amount of chitin in S. monoica accounts for less than 0.5% of the total cell wall carbohydrates (Bulone et al., 1992). Guerriero et al. (2010) have shown, by using nikkomycin Z as a specific inhibitor of chitin synthase, that this enzyme is most likely involved in producing chitin at the tip of S. monoica hyphae as tip bursting was observed in the presence of the drug.

Recent analytical work by Mélida et al. (2013) on oomycete mycelial walls led to a new classification of Peronosporales and Saprolegniales into three groups based on their cell wall structures. The work was performed on 10 different species. Table I summarizes the composition of each cell wall type.

Table 1. Criteria characterizing the three major types of oomycete cell walls according to Mélida et al. (2013).

The Peronosporales are representative of type I cell walls (Mélida et al., 2013). Two species were studied, Phytophthora parasitica and P. infestans. Type I cell walls are devoid of GlcNAc, and exhibit the highest content in mannose (>3% of the cell wall content) compared to the two other types of cell walls (<3%). The alkali soluble fraction of type I cell walls contains glucuronic acid (GlcA), as opposed to type II and III cell walls (Table I). Until recently, the quantity of cellulose in the genus Phytophthora was thought to be around 15% (Aronson et al., 1967). In fact it was estimated that Phytophthora species contain between 32-35% of cellulose based on the assumption that all 1,4-linked glucosyl residues arise from cellulose (no starch-

8 like polysaccharide are found in Phytophthora) (Mélida et al., 2013). This discrepancy is probably due to the techniques used that were different between the earlier and more recent study (Mélida et al., 2013). Cellulose can occur as a crystalline or amorphous polymer. The GC/MS technique used by Mélida et al. (2013) cannot distinguish these different forms of cellulose and provides a quantification of all (1,4)- linked glucans, whereas the X-ray diffraction analysis used in Aronson’s work (1967) detects crystalline cellulose only.

The cell walls of four genera of Saprolegniales (Achlya, Dictyuchus, Leptolegnia, and Saprolegnia) were classified as type II. Unlike type I cell wall, GlcNAc is present in their cell walls and the content is lower than 5% and they are devoid of GlcA. It is the only type of cell walls where 1,3,4-linked glucosyl residues was indicated to crosslink cellulose and 1,3-glucans and these residues were found in an alkali-insoluble fraction (Mélida et al., 2013) (Table I). Type II cell walls contain the highest amount of cellulose of all cell wall types (more than 40%).

Aphanomyces was the only oomycete genus from the different organisms studied that contains type III cell walls. The latter has a GlcNAc content higher than 5% and they are the only type of cell walls in which unusual 1,6-GlcNAc residues occur (Mélida et al., 2013) (Table I). It is the first time that this type of monomer is described in eukaryotic cell wall polysaccharides. The occurrence of 1,6-GlcNAc- based carbohydrates has however been reported in biofilms formed by Escherichia coli and Staphylococcus epidermidis (Götz, 2002; Itoh et al., 2005). The recent study by Mélida et al., (2013) contradicts previous works that did not detect chitin but detected other GlcNAc-based carbohydrates (10%) in the cell wall of A. euteiches (Badreddine et al., 2008).

1.3.2 Fungal and yeast cell walls

Fungal and yeast cell walls are more well-studied than oomycete cell walls and thus, a background of cell wall compositions from these organisms will be provided. As opposed to oomycetes and plants, fungi and yeasts do not contain cellulose (Figure 5). Cell wall components and structure differ between fungal species, developmental

9 stages and varying growth conditions. Fungal cell walls consist mainly of β-glucans, chitin, mannan and glycoproteins usually found in the form of highly glycosylated mannoproteins (Bowman and Free, 2006). Similar to oomycetes, glucans are the major cell wall structural polysaccharides making up 50-60% of the dry weight of fungal cell walls. β-1,3-Glucans and chitin form a cross-linked network and are considered the scaffolding structure of the cell wall.

Due to these polysaccharides importance in maintaining the structural integrity of their cell walls, β-1,3-glucans and chitin represent an interesting target for disease control. Antifungal drugs targeting fungal cell wall have been found to be useful for fungal control. For example, Echinocandins a drug that inhibits β-1,3-glucan synthase, has been found to be promising in providing cure for aspergillosis and candidiasis (Odds et al., 2003). Several other inhibitors, such as anidulafungin, caspofungin and micafungin have also been shown to interfere with the biosynthetic process of β-1,3-glucans (Odds et al., 2003). The highest level of growth inhibition of Candida albicans and Aspergillus fumigatus is obtained by using a combination of chitin synthase and glucan synthase inhibitors (Walker et al., 2008; Verwer et al., 2012). The catalytic subunits of the β-1,3-glucan synthases of Saccharomyces cerevisiae are considered to be encoded by two genes, Fks1 and Fks2 (Douglas et al., 1994; Mazur et al., 1995), although this has not been unequivocally demonstrated. Mutations in Fks1 slow down the growth of yeast cells and alter cell wall structure, while simultaneous mutations in both genes are lethal for the organism (Douglas et al., 1994; Mazur et al., 1995).

Chitin β-1,3-glucans CW β-1,6-glucans Mannans PM Proteins

GPI

Figure 5. Cell wall model of C. albicans (adapted from Gow, 2012).

Fungal and yeast β-glucans can be divided into different subtypes based on the type of linkages they contain. β-1,3-Glucans are typically long polymers of around 1500

10 glucose residues. They account for 85% of the total yeast cell wall β-glucans (Manners et al., 1973). Another subtype, β-1,6-glucans, typically consists of short chains of up to 300 glucose residues. This subtype represents about 15% of the total yeast cell wall β-glucans (Klis et al., 2002) and is involved in cross-linking different β- glucan polymers (Cabib et al., 2001). β-1,3-Glucans may also contain β-1,6 branches, which gives rise to β-(1,3;1,6)-glucans, typical of yeast and fungal cell walls. β-1,6-Glucans are noncrystalline and is believed to interact with β-1,3-glucans, mannoproteins and chitin (Klis et al., 2002; Kollár et al., 1997). To date, linear β-1,6- glucans have not been shown to occur in filamentous fungi such as Neurospora crassa and A. fumigatus (Borkovich et al., 2004; Fontaine et al., 2000). However, this is not to suggest that other fungal species do not synthesize linear β-1,6-glucans.

Another key polysaccharide of fungal and yeast cell walls is chitin. Chitin is a minor constituent of S. cerevisiae cell walls and accounts for only 1 to 2% of the cell wall dry weight. However, the presence of chitin is required for the septation process (Orlean, 2012). Chitin content is higher in some fungal species, including Aspergillus and Neurospora sp., where it may make up to 20% of the cell wall dry weight (Bartnicki-Garcia, 1968). Chitin chains assemble via hydrogen bonds to form microfibrillar and crystalline structures with high tensile strength, contributing to the strength and integrity of the cell wall. The inhibition of chitin synthases results in disordered cell walls, cell deformation and reduced control of osmolarity (Specht et al., 1996).

In addition to polysaccharides, glycoproteins represent an important fraction of the cell wall. In the yeast species S. cerevisiae and C. albicans, glycoproteins make up approximately 30 to 50% of the cell wall dry weight, whereas filamentous fungi have a slightly less dry cell wall glycoprotein content of 20 to 30% (Brown and Catley, 1992). Glycoproteins interact with chitin and the matrix of glucans (Marcilla, 1991) and, they are essentially represented by mannoproteins in the yeast such as S. cerevisiae and C. albicans. For filamentous fungi, in addition to mannoproteins, galactose residues also occur in the glycan structures of the cell wall glycoproteins (Goto, 2007; Lommel and Strahl, 2009). The biosynthesis of glycoproteins which involves the glycosylation process starts in the lumen of the endoplasmic reticulum

11 and that proceed further in the Golgi apparatus where glycans maturation take place (Lesage and Bussey, 2006).

In both yeast and fungi, glycosylphosphatidylinositol (GPI) anchored proteins represent a fraction of the wall-bound proteins (Fontaine et al., 2003). Figure 6 represents the core structure of the GPI anchor protein that is made of: a phosphatidylinositol, a core tetrasaccharide (composed of 3 mannose residues and one glucosaminyl residue), and a phosphoethanolamine. The protein is linked to the phosphoethanolamine group by forming a bond between its COOH terminus and the ethanolamine group of the anchor. Some GPI proteins are enzymes, for instance, 1,3-β-glucanosyltransferases is involved in the synthesis of glucans (Mouyna et al., 2000) and glycosidases (Rodriguez-Peña et al., 2000). They contribute to the reorganization of the cell wall, and belong to the GPI protein family. Studies on the biosynthesis of GPI anchors in S. cerevisiae demonstrated their key role in cell wall integrity (Leidich et al., 1994; Kawagoe et al., 1996 and reviewed in Orlean, 2012). GPI proteins do not occur in bacteria (Eisenhaber et al., 2001) with the exception of an ancient group of Archaea bacteria (Kobayashi et al., 1997) and enzymes involved in the biosynthesis of GPI anchors differ between mammalians and parasites like Trypanosoma brucei (Nagamune et al., 2000) and Plasmodium falciparum (Naik et al., 2000). These differences suggest the biosynthetic pathways of GPI anchors in different groups of organisms represent targets for the development of specific antiparasitic agents (Doering et al., 1991; Leidich et al., 1994).

Figure 6. The GPI anchor structure. The three principal units are the phosphatidylinositol, the core tetrasaccharide and the phosphoethanolamine. R1 and R2 represent the fatty acid residues.

1.4 Carbohydrate-Active enZymes (CAZymes)

As previously mentioned, enzymes involved in cell wall biosynthesis or carbohydrate- active enzymes (CAZymes) can be exploited for disease control by identifying substances that block these enzymes. In particular, CAZYmes from GT2 family will be described as two key enzyme domains studied in this thesis belong to enzymes from this family.

In CAZy, the carbohydrate-active enzyme database (www.cazy.org., Campbell et al., 1997), carbohydrate-active enzymes (e.g., glycosyltransferases (GTs), glycoside hydrolases (GHs)) are classified based on amino acid sequence similarities and, when available, structural fold. Chitin synthases are classified in the GT2 family together with cellulose synthases. As of September 2015, there are 97 different GT

13 families described in the CAZy database. GT2 enzymes studied to-date adopt a GT- A structural fold (Coutinho et al., 2003; Lairson et al., 2008). It should be noted, however, that a large proportion of GT2 members have yet to be structurally characterized. Thus, it is possible that some members of the GT2 family may adopt alternative folds. In addition to sequence and structural similarities, all enzymes in a given family share the same catalytic mechanism (Henrissat et al., 1997).

GTs form glycosidic bonds by catalyzing the transfer of sugars from activated sugar donors (e.g., UDP-GlcNAc for chitin synthase and UDP-Glc for cellulose synthase) to specific acceptor chains (Figure 7). Chitin synthases (Chs) and cellulose synthases (CesA) are membrane-associated inverting GTs with N-acetylglucosaminyl transferase activity (EC 2.4.1.16) and glucosyltransferase activity (EC 2.4.1.12), respectively. A high proportion of GTs are involved in cell wall biosynthesis by forming different types of glycosidic linkages in carbohydrate polymers. It has been postulated that there are at least as many GTs as the types of linkages formed. Some GTs, however, are able to form several types of linkages. It is the case for instance of hyaluronan synthase, which forms an unbranched polysaccharide composed of repeating disaccharide units of GlcNAc and Glc A linked via alternating β-1,4 and β-1,3 glycosidic bonds.

Figure 7. Reaction catalysed by chitin synthases.

GTs can synthesize glycan chains through two types of mechanisms: inverting or retaining. The stereochemistry with respect to the nucleotide-sugar (α configuration) either changes (inverting mechanism) or is conserved (retaining mechanism) during catalysis. GT2 family members (e.g. CesA, Chs) have an inverting mechanism including a one-step reaction in which a β-linked product is formed from the α-linked UDP-sugar donor (Figure 8).

Figure 8. Proposed mechanism for inverting glycosyltransferases (after Charnock et al., 2001).

The reaction involves two acidic residues of the enzyme active site. A first caboxylate acts as a base, deprotonating a hydroxyl group of the acceptor sugar, thereby activating it (Charnock et al., 2001). This activation allows the nucleophilic attack by the acceptor molecule of the anomeric carbon of the nucleotide-sugar donor. The carboxylic group of the second catalytic amino acid then facilitates the departure of the nucleotide leaving group whose phosphate groups are stabilized by coordination with a divalent cation (Mg2+, Mn2+) (Charnock and Davies, 1999; Charnock et al., 2001) (Figure 8). Retaining enzymes use a two-step mechanism which involves the formation of a covalent glycosyl-enzyme intermediate (Koshland, 1953). One catalytic residue acts as a general acid/base. The second catalytic residue acts as a nucleophile, attacking the glycosyl-enzyme intermediate during the second step of the reaction and leading to the release of the glycosylated product whose configuration is identical to that of the sugar donor (Figure 9).

Figure 9. Proposed mechanism for retaining glycosyltransferases (adapted from Yip and Withers, 2004). Divalent cations (M2+) play similar role as in the inverting mechanism. Abbreviations: R, a nucleotide; R’OH, an acceptor group for instance another sugar or a protein.

As discussed previously, the product of GTs such as chitin, is part of the oomycete and fungi cell wall content. The next section will discuss about the structure of chitin and its different allomorphs found in nature.

1.5 Chitin

1.5.1 Chitin structure

Chitin is a linear polymer that can contain over 1000 GlcNAc residues linked by β- (1,4)-linkages. It is not soluble in aqueous media due to its high molecular weight and crystalline structure. Chitin is polymorphic, as it occurs as three different allomorphs designated α, β and γ (Figure 10). The allomorphs can be distinguished by X-ray diffraction analysis (Rudall and Kenchington 1973; Kramer and Koga 1986; Imai et al., 2003). The α structure is by far the most prevalent in nature, for instance in the

16 exoskeletons of arthropods, crab shells and cell walls of fungi, α-chitin serve as the load bearing component (Cohen, 1993). α-Chitin consists of an antiparallel arrangement of chitin chains which are tightly packed and held as a coherent and crystalline structure through multiple intermolecular and intramolecular hydrogen bonds and van der Waals forces that altogether provide rigidity (Minke and Blackwell, 1978). In the antiparallel structure of α-chitin, the reducing ends of the consecutive chains in the same sheet point to opposite directions. Figure 11 shows the intramolecular hydrogen bonds occuring between the hydroxyl group (OH) at C-6 of one sugar residue and the carbonyl group (C=O) of the adjacent monosaccharide. Intramolecular hydrogen bonds are also present between the hydroxyl group at C-3 and the oxygen atom in the ring of an adjacent residue.

Figure 10. Representation of the different allomorphs of chitin. α-Chitin is characterized by antiparallel chains whereas β-chitin consists of parallel chains. γ-Chitin consists of an alternative succession of one antiparallel and two parallel chains.

Figure 11. Chitin structure and intramolecular hydrogen bonds (dotted lines).

The β structure is less common (<1% of the chitin found in nature) and consists of parallel chains. Compared to α-chitin, β-chitin contains fewer interchain hydrogen

17 bonds and is therefore less rigid and can swell more easily in water (Blackwell, 1969). Highly crystalline β-chitin can be found for example in the pogonophore tubes (Tevnia jerichonana) (Gaill et al., 1992) and squid pens (Loligo sp). These structures are the richest sources of β-chitin in nature (Lavall et al., 2007).

The third allomorph, γ-chitin, was proposed by Rudall (1963), but its existence remains controversial. This allomorph would consist of a succession of two parallel chains followed by a single antiparallel chain (Figure 10). The larvae of some insects such as silkworm (Antheraea pernyi) and sawfly (Phymatocera aterrima) have been described to contain some γ-chitin (Rudall and Kenchington, 1973). β- and γ-chitins are more flexible and softer than α-chitin. Indeed, the packing of individual chains is less tight in these allomorphs due to a lower number of inter-chain hydrogen bonds and a higher interaction with water. These two types of chitin can be converted to α- chitin by chemical treatments. The incubation of γ-chitin in a saturated solution of lithium thiocyanate at room temperature leads to the formation of α-chitin. The least crystalline forms of β-chitin (e.g., from Loligo pen) can be converted to α-chitin by treatment with 6 N hydrochloric acid solution at cold temperatures (Rudall and Kenchington, 1973).

The different types of allomorphs presented above are produced by chitin synthases found in different organisms. In section 1.6 the biosynthesis of chitin, the chitin synthases and their post-translational regulation as well as their transport to the plasma membrane will be discussed.

1.6 Chitin biosynthesis

1.6.1 Main process

The detailed molecular mechanisms responsible for chitin biosynthesis are not well understood (Cohen, 2001). In fact, many of the same questions remain unanswered for both chitin and cellulose biosynthesis. While the mechanism of cellulose chain translocation across the plasma membrane (PM) has been elucidated recently in the

18 bacterium Rhodobacter (Morgan et al., 2013), the mode of transport of chitin across the membrane remains elusive. It is however likely that the catalytic subunits themselves are able to extrude individual chains as is the case for cellulose biosynthesis in Rhodobacter (Morgan et al., 2013). The mode of assembly of individual chains into microfibrillar structures is yet another unknown process for both chitin and cellulose biosynthesis. Despite decades of efforts to decipher these processes, to date no potential protein that may be involved in chain assembly has been identified. This suggests that microfibril assembly is essentially a spontaneous process.

Chs genes have been identified in many organisms, including fungi and oomycetes, where chitin is typically synthesized in regions of active growth (e.g. hyphal tip) and cell wall remodelling. However, the mode of interaction of chitin with the other cell wall components as well as the molecular events responsible for the establishment of these interactions remain unknown. In yeast, chitin is primarily synthesized at the bud tip during growth and at the bud neck during cytokinesis (Cabib et al., 2001). In filamentous fungi, chitin is essentially synthesized at the hyphal apex. This was indirectly demonstrated in Mucor rouxii where it was shown that the Chs enzymes are essentially localized at the hyphal tip (McMurrough et al., 1971). In this example, the site of chitin biosynthesis was demonstrated by autoradiography using a radioactive substrate of chitin synthase, UDP-[3H]GlcNAc (McMurrough et al., 1971). Before Chs can perform chain elongation, the substrate UDP-GlcNAc must be synthesized. This involves several consecutive enzymatic reactions (Figure 12).

20 Figure 12. The chitin synthesis pathway. In fungi, glucose is the starting molecule that is modified through several consecutive steps to achieve synthesis of UDP-N-acetylglucosamine. The enzymatic reactions leading to the synthesis of UDP-N-acetylglucosamine occur in the cytoplasm. The enzymes producing chitin is located in the plasma membrane. This representation is based on previously published pathways (Kramer and Koga, 1986; Cohen, 2001 and Merzendorfer and Zimoch, 2003).

1.6.2 Chitin synthases

Chitin synthases have been well studied in the yeast S. cerevisiae. Three Chs were isolated from this species (Bulawa et al., 1986; Sburlati and Cabib 1986; Valdivieso et al., 1991). Chs1 appears to be involved in cell wall repair after excessive action of chitinase at the bud scar (Cabib et al., 1989). Chs2 is required during cytokinesis, for the formation of the primary septum between the mother and daughter cells (Silverman et al., 1988). Chs3 is involved in the biosynthesis of 80 to 90% of the total amount of chitin in the cell wall (Shaw et al., 1991). Mutating Chs2 and Chs3 genes is lethal for S. cerevisiae (Shaw et al., 1991). As opposed to yeast, most fungal species contain a higher number of Chs genes, typically between 3 to 6 (Mellado et al., 1995). The genome of some organisms has been shown to contain even more genes, e.g. 8 in Aspergillus nidulans and Ustilago maydis, and, up to 10 in Phycomyces blakesleeanus (Horiuchi, 2009; Weber et al., 2006; Miyazaki and Ootaki 1997). The multiple number of Chs genes could be related to the diversity of the morphologies found in filamentous fungi (Chigira et al., 2002), although some Chs genes may also have redundant functions. The multiplicity of Chs genes could provide the organism the opportunity to adapt to changes in the environment, thereby facilitating growth in different ecological niches (Ruiz-Herrera et al., 2002). Other studies based on cell localization and mutations in U. maydis have suggested that multiple Chs isoforms cooperate (Weber et al., 2006). Experiments have shown that the mutation of a single Chs gene often does not show any phenotype, whereas double or triple mutants show a phenotype (Specht et al., 1996; Wang et al., 2001). The oomycete Chs genes have been less studied than their yeast and fungal counterparts. However, research on oomycete Chs has gained interest during the past years as these enzymes represent potential targets for disease control (Guerriero et al., 2010). Until recently, oomycete species were considered to contain two Chs genes. However the sequencing of the S. parasitica genome has revealed

21 the presence of 6 putative Chs genes in this organism (Jiang et al., 2013). It should be emphasized however that, with the exception of the chitin synthase 2 from S. monoica (SmChs2), the activity of other gene products has not been demonstrated biochemically (Guerriero et al., 2010).

Figure 13 shows the predicted domain organization of the oomycete chitin synthases after translation of the corresponding genes to their amino acid sequences. The number of genes can vary within the same genus, as illustrated by the S. monoica (2 genes) and S. parasitica (6 genes) species. SmChs2 has been characterized biochemically using a recombinant form of the enzyme, whereas the same approach failed to demonstrate any chitin synthase activity in vitro for SmChs1 (Guerriero et al., 2010). Despite chitin content in the corresponding hyphal walls is typically low (no more than 1-2%), chs genes are found in most of the oomycete species studied to- date, and, chitin was found to be vital for cell survival as demonstrated in S. monoica (Guerriero et al., 2010). Interestingly, the Saprolegniales Aphanomyces euteiches contains 2 putative Chs genes and cell wall analysis of the mycelium showed the occurrence of around 10% of GlcNAc-based carbohydrates (Badreddine et al., 2008). A small fraction of these carbohydrates seems to correspond to true chitin, and the rest being soluble chitooligosaccharides (Mélida et al., 2013). These observations suggest that Chs genes may be responsible for the biosynthesis of different types of GlcNAc chains. Most intriguingly, the genome of the Peronosporales P. infestans contains only one putative Chs gene (Mort-Bontemps et al., 1997), but no chitin is detectable in the cell wall of the mycelium (Mélida et al., 2013). Based on these observations, it has been proposed that P. infestans putative Chs gene may be involved in processes other than chitin biosynthesis, including the formation of GPI- anchored proteins (Mélida et al., 2013).

Chitin synthases are membrane-bound proteins that usually contain seven transmembrane domains (TMDs) (Figure 13). Most of these domains are located at the C-terminal ends of the proteins (Figure 13). In many fungal Chs, a cluster of 3 to 5 TMDs follows the catalytic domain. Enzymes with a cluster of 5 TMDs show the presence of 2 other TMDs at their C-terminal. For instance the oomycete and yeast Chs shown in Figure 13 are following this type of organization.

22 Some oomycete Chs contain a domain with unknown function similar to the so-called “Microtubule Interacting and Trafficking domains” (MIT) found in other organisms. (Figure 13 and Guerriero et al., 2010; Rzeszutek et al., 2015, manuscript).

Figure 13. Domain comparison of oomycete Chs (the diagram was made using DomainDraw (Fink and Hamilton, 2007)). Chs proteins from oomycetes share a similar transmembrane domain pattern. The MIT domain (yellow) is a feature that differentiates SpChs1, SpChs2, SmChs1 and SmChs2 from the other oomycete Chs. Manual sequence analyses suggest the occurrence of an N-terminal MIT domain in AeChs1, which was not identified by bioinformatics searches as a typical Pfam domain. MMD (Myosin Motor-like Domain) is present only in some fungal Chs. (adapted from Rzeszutek et al., 2015, manuscript).

24 As for most processive GTs, Chs contain the two conserved motifs D,D,D and QXXRW (Figure 13). Structural analysis of SpsA, a bacterial GT2 enzyme of unknown function, suggests that these motifs form a single center for glycosyl transfer (Charnock et al., 2001). Sequence analysis of proteins with demonstrated function and catalytic activity have shown that, except for some cases, domains A (D,DXD) and B (D,QXXRW) are present in processive GTs while non-processive enzymes contain only domain A (Saxena et al., 1995). The crystal structure recently obtained from the cellulose synthase of Rhodobacter sphaeroides (Morgan et al., 2013) has provided insights into the function of the two previously described conserved motifs (Charnock et al., 2001; Saxena et al., 2001). The first two aspartic acids bind the nucleotide-sugar while the third aspartic acid is believed to be the catalytic base. The latter residue is indeed in close contact with the non-reducing end of the glucan acceptor. A binding site comprising the QXXRW motif and the conserved FFCGS sequence is responsible for holding the growing acceptor chain in the active site of the enzyme (Morgan et al., 2013). Chs are processive enzymes, like cellulose synthases. The chitin chain remains bound to the enzyme during polymerization, allowing the successive addition of multiple GlcNAc residues at the non-reducing end of the elongating polymer (Charnock et al., 2001; Imai et al, 2003). A similar mechanism (elongation from the non-reducing end) was experimentally demonstrated for the polymerisation of cellulose chains in the bacterium Gluconacetobacter xylinus (Koyama et al., 1997) and the plant Rubus fruticosus (blackberry) (Lai Kee Him et al., 2002). The importance of the D,D,D and QXXRW motifs for Chs has been studied in yeast by site-directed mutagenesis. Mutations in either of the motifs lead to a significant decrease in Chs 2 activity in S. cerevisiae (Nagahashi et al., 1995). Similar results were obtained with point mutations in the QXXRW motif of S. cerevisiae Chs3 (Cos et al., 1998). Mutations in these motifs in the G. xylinus CesA also show their importance for enzyme activity (Saxena and Brown, 1997; Saxena et al., 2001).

25 1.6.3 Post-translational regulation of chitin synthases

1.6.3.1 Phosphorylation

Phosphorylation plays an important role in determining the location and stability of Chs. For instance, in S. cerevisiae Chs2, the four phosphorylation sites located at the N-terminus of Chs led to the retention of enzyme in the ER (Teh et al., 2009). Deletion of the corresponding phosphorylation sites also resulted in the degradation of Chs2 (Martinez-Rucobo et al., 2009). Moreover, the proper targeting of Chs3 in C. albicans cells have been shown to require a phosphorylation event (Beltrao et al., 2009), thus, Chs transport and integrity seemed to be both regulated and controlled by phosphorylation.

1.6.3.2 Zymogenicity

Cabib and Farkas (1971) were first to show that the Chs of S. cerevisiae and Saccharomyces carlsbergensis must be cleaved by proteases prior to becoming catalytically active. Trypsin has been shown to activate Chs in vitro and this is the case for Chs from e.g. M. rouxii (McMurrough and Bartnicki-Garcia, 1973; Ruiz- Herrera and Bartnicki-Garcia 1976), Aspergillus flavus, Aspergillus nidulans (Lopez- Romero and Ruiz Herrera 1976; Ryder and Peberdy, 1977) and the genera Agaricus, Allomyces and Neurospora (Bartnicki-Garcia et al., 1978). In S. monoica, Chs2 was also found to be more active in the presence of trypsin (Guerriero et al., 2010). Recently, Chs activity in microsomal preparations from A. euteiches was shown to be higher when trypsin is present in the reaction mixture (Bottin et al., 2015 manuscript). Keeping Chs in an inactive zymogenic form may be a way for the cell to regulate chitin biosynthesis in a spatio-temporal manner. Whether trypsin acts directly on the Chs enzymes or indirectly by activating other proteins remains unknown. However, experiments on the zymogenicity of S. cerevisiae Chs2 showed that trypsin is acting indirectly on an unknown soluble protease, resulting in a stimulation of Chs2 activity (Martínez-Rucobo et al., 2009).

26 1.6.4 Targeting of chitin synthases to the plasma membrane

1.6.4.1 Chitosomes

Chitin synthases are associated to the plasma membrane as has been revealed through pioneering experiments using protoplasts of S. cerevisiae (Durán et al., 1975) and Schizophyllum commune (Vermeulen et al., 1979). However, the processes by which Chs is targeted to and inserted in the plasma membrane remain unknown, especially in oomycetes.

Small spheroidal vesicles (40-70 nm) called chitosomes were first isolated from the fungus M. rouxii and shown to be able to synthesize chitin in vitro (Bracker et al., 1976). To confirm the broader occurrence of chitosomes across different fungal organisms, their isolation was conducted in Allomyces macrogynus (Chytridiomycetes), N. crassa (Ascomycetes), S. cerevisiae (Ascomycetes), Agaricus bisporus (Basidiomycetes) and M. rouxii (Zygomycetes) (Bartnicki-Garcia et al., 1978). No differences in chitosome structure and size were found between these species. However, the number of chitosomes and the stability of Chs varied depending on the life stages of the fungus. In most cases, a fibrillar material could be visualized inside the chitosome preparations (Bartnicki-Garcia et al., 1978).

Despite these data, the existence of chitosomes was debated for some time (Duran et al., 1979; Farkas, 1979). Some researchers suggested that chitosomes are endocytosis products or artefactual structures that form during cell disruption. The argument that chitosomes are a product of endocytosis was eventually ruled out based on many observations. In particular, chitosomes exhibit a lipid and protein composition that differs from the plasma membrane (PM) (Bracker et al., 1976; Flores-Martinez et al., 1990) and their membrane is thinner than that of the PM, vacuolar and secretory vesicles (Bracker et al., 1976). The improvement of centrifugation procedures required to isolate chitosomes has allowed the separation of two populations of Chs (Leal-Morales et al., 1988). Another characteristic of the yeast plasma membrane is the presence of β-(1-3)-glucan synthases, which have never been reported in chitosome preparations (Leal-Morales et al., 1988). Another indication that chitosomes are not a product of endocytosis is the observation that

27 Chs from chitosomes are catalytically inactive in vivo whereas the plasma membrane enzymes are active. Altogether, these data support the theory that chitosomes are unique intracellular compartments which, deliver Chs to the cell surface (Bracker et al., 1976; Bartnicki-Garcia, 2006).

1.6.4.2 Myosin motor-like domains of Chs

Myosin motor-like domains (MMD) are located at the N-terminal of some chitin synthases from filamentous fungi, dimorphic fungi and bivalves (Weiss et al., 2006; Treitschke et al., 2010). For instance, one of the Chs from the dimorphic fungus Ustilago maydis (UmMcs1) has an MMD (Weber et al., 2006). Similarly, two of the eight Chs from Aspergillus nidulans have an MMD, and were designated CsmA and CsmB, respectively for chitin synthase with a myosin motor-like domain (Fujiwara et al., 1997; Takeshita et al., 2006). MMDs vary in length depending on whether they contain conserved ATP-binding motifs (P-loop, Switch I and II) thought to be important for ATPase and motor activities (Takeshita et al., 2006). Orthologues of CsmA and CsmB have been found in other filamentous and dimorphic fungal species but not in S. cerevisiae, Schizosaccharomyces pombe or oomycetes. Phylogenetic analyses classify the unconventional myosin class XVII as specific to Chs (Hodge and Cope, 2000).

Previous in vitro co-sedimentation studies on CsmA and CsmB have shown that there are interactions between MMDs from these proteins and F-actin (Takeshita et al., 2005; Takeshita et al., 2006). Interestingly, actin plays a key role in both tip growth and septum formation, i.e. at sites where chitin is synthesized and is more concentrated at the hyphal tip during periods of growth. It has been suggested that Chs may be transported to the exocytosis sites via MMD (Fujiwara et al., 1997; Madrid et al., 2003). Studies have shown that the MMD of CsmA and CsmB is essential for the proper localization and function of the enzyme (Takeshita et al., 2005; Takeshita et al., 2006).

Besides actin, MMD was shown to also interact with microtubules. Recent microscopy studies have shown the long range motility of UmMcs1 along MTs (Treitschke et al., 2010; Schuster et al., 2012) and the motility of UmMcs1 was

28 affected in the presence of Benomyl, a drug that disrupts MTs (Schuster et al., 2012). In the same study, vesicles were found to be transported along the MTs by kinesin-1 and along the peripheral of F-actin by myosin-5. A truncated UmMcs1 protein devoid in MMD shows a lower exocytosis rate than the full-length protein (Treitschke et al., 2010).

Both F-actin and microtubules (MTs) together with MMD, appear to be important in fungal tip growth although their precise functions remain unknown (Taheri-Talesh et al., 2008). Vesicles containing Chs may use the MTs and F-actin tracks to accumulate at the apical region of the hyphae to form Spitzenkörpers. Spitzenkörpers act as “vesicle supply centres” (VSC) to respond to the delivery needs at the hyphal apex and are believed to be essential for hyphal tip extension (Bartnicki-Garcia et al., 1989). Although Spitzenkörpers are assumed to contain Golgi vesicles carrying secretory proteins and cell wall precursors, this theory has yet to be supported by experimental data (Steinberg, 2007).

It is noteworthy that transport of vesicles to the apex does not guarantee their insertion to the PM. In fact, only 15% of vesicles fused successfully to the PM (Schuster et al., 2012) while the remaining are recycled via interactions with dynein, yet another motor protein (Schuster et al., 2012). Similar proportions of fusion events were found in animal cells (Toonen et al., 2006). Successful fusion is dependent on the residence time of the vesicle by the plasma membrane. In animal cells, successful exocytosis is defined by an extended tethering time (>10 s) at the plasma membrane (Toonen et al., 2006). The residence time is essential not only for the successful insertion of Chs in the plasma membrane, but also for the initial phase of pathogenicity as has been shown in Fusarium oxysporum (Madrid et al., 2003) and U. maydis (Weber et al., 2006).

In conclusion, altogether these data suggest that MMDs may assist the secretion of Chs via interactions with actin and/or microtubules

29 1.6.4.3 MIT domains

The oomycete S. monoica does not contain any chitosome (Leal-Morales et al., 1997). Therefore a different mechanism must be employed by oomycetes to transport their Chs to the plasma membrane. In contrast to fungi, some oomycete Chs contain a so-called Microtubule Interacting and Trafficking (MIT) domain at their N-terminal end (Figure 13). This has been demonstrated in Chs from S. monoica (Brown et al., submitted). Putative MIT domains are also present in other oomycete species, including S. parasitica. The function of the MIT domain in carbohydrate synthases is unknown, but it may be involved in delivery or/and recycling/degradation processes (Brown et al., submitted). More detailed discussions on MIT domains and their function are provided in section 1.9 and 1.10.

1.7 Cellulose

One of the main differences between oomycete and fungi is the presence of cellulose in the cell wall of oomycete. To better understand cellulose biosynthesis in oomycete, in addition to what is known in oomycete, plant cellulose biosynthesis will also be described in the next section. Similar unknown questions regarding the biosynthesis can be addressed in both organisms.

Cellulose is responsible for the strength and rigidity of plant cell walls (Brown, 1996) and parts of the exoskeleton of urochordates (e.g., ascidians) (Matthysse et al., 2004). In oomycetes, cellulose is thought to play a similar role to that of chitin in true fungi by contributing significantly to the strength of the cell wall (Farkas, 1979). Cellulose is a linear homopolymer of glucose residues linked by β-(1,4) linkages. Cellobiose is sometimes considered the repeating unit of cellulose because every glucose unit of the cellulose chain is rotated by 180º with respect to its neighbour (Figure 4b). The strength of cellulose is due to the extended organization of the linear chains, which favours a tight packing of the individual molecules into stable microfibrils. Similar to chitin, cellulose is stabilized by intramolecular and intermolecular hydrogen bonds, leading to the formation of crystalline structures.

30 1.7.1 Terminal complexes

Cellulose synthase complexes were first observed in the plasma membrane by freeze-fracture at the tip of the growing cellulose microfibrils of the alga Oocystis apiculata (Brown and Montezinos, 1976). For this reason, they were designated as terminal complexes (TCs). Plant TCs exhibit hexagonal supramolecular structures designated “rosettes” due to their six-fold symmetry (Mueller and Brown 1980; Brown 1996).

A “rosette” consists of six subunits, each of which is predicted to contain 6 catalytic cellulose synthase (CesA) proteins that correspond to the products of three different CesA genes (Doblin et al., 2002). Based on this organization, a rosette would consist of 36 CesA proteins and form microfibrils that would contain 36 glucan chains (Figure 14). However, measurements using Wide-Angle X-ray Scattering and Solid-State NMR on mung bean (Vigna radiata) primary cell walls suggest that the number of glucan chains formed would be rather 18 or 24 instead of 36 cellulose chains (review in Guerriero et al., 2010; Bootten et al., 2004; Newman et al., 2013). This may imply that there would be less than 6 CesAs per subunit or that not all of the CesAs in the subunit are active.

Other organisms such as algae and some bacteria produce cellulose, but the corresponding biosynthetic enzymes are not organized as rosettes as in plants. Instead, the complexes are arranged as arrays of multiple catalytic subunits designated linear TCs (Brown, 1996). Cellulose also occurs in the cell walls of oomycetes, but to-date no terminal complexes have been identified in these organisms.

1.7.2 Cellulose structure and biosynthesis

The natural form of cellulose is called cellulose I or native cellulose. In these microfibrils, the glucan chains are organized in a parallel fashion (Koyama et al.,

1997). Cellulose I can occur as two different allomorphs designated Iα and Iβ (Atalla and VanderHart 1984). These two allomorphs co-exist in plant cell walls whereas other sources of cellulose are much richer or contain exclusively one of the two

31 allomorphs. This is the case of bacterial cellulose and some algal cellulose e.g.

Valonia, which are typically enriched in the Iα allomorph (Sugiyama et al., 1991) whereas in tunicates, they contain only the Iβ allomorph (Belton et al., 1989).

Cellulose IVI is also a natural form of cellulose. It corresponds to a disorganized form of cellulose I and typically occurs in the primary cell walls of plant cell suspension cultures and in oomycetes (Bulone et al., 1992; Helbert et al., 1997). Cellulose II and III can be prepared by various chemical treatments, but they are not encountered in nature. Cellulose II, as opposed to cellulose I, consists of antiparallel chains, and is widely exploited in the textile industry (viscose).

Despite many years of research, the molecular mechanisms of cellulose biosynthesis remain poorly understood. There is a great interest in improving our understanding of the cellulose biosynthesis process as this knowledge can be exploited for many applications, e.g. the development of herbicides, bioethanol production from plant biomass and the development of green materials.

Most of the knowledge we do have about cellulose biosynthesis comes from studies in bacteria and in higher plants. These studies show cellulose biosynthesis is a complex process involving several steps. Figure 14 shows a hypothetical model proposed for cellulose biosynthesis in plants and highlights the outstanding questions corresponding to the different steps implicated in the cellulose biosynthesis process (Guerriero et al., 2010).

Figure 14. Hypothetical model describing cellulose biosynthesis (adapted from Guerriero et al., 2010). The model shows the formation and extrusion of a glucan chain from a CesA catalytic subunit through the plasma membrane. Many aspects of cellulose biosynthesis remain unknown and are indicated in this model by question marks.

This model for plant cellulose biosynthesis can now be revised by integrating data from the recently solved structure of the bacterial cellulose synthase (BcsA) in the presence of a translocating glucan chain and a UDP-glucose molecule (Morgan et al., 2013). This first structure of a cellulose synthase has been exploited to model the cellulose synthase from cotton (Gossypium hirsutum) (Sethaphong et al., 2013; Slabaugh et al., 2013). From the crystal structure, it is possible to identify a channel that forms from the third to the eighth transmembrane domain of the protein and that allows the translocation of the glucan chain across the plasma membrane (Morgan et al., 2013).

UDP-Glucose (UDP-Glc), the substrate of CesA, can be synthesized by both UDP- Glc pyrophosphorylase, a cytosoluble enzyme, and by sucrose synthase (SuSy). A membrane-bound form of SuSy was described in developing cotton fibers and it has

33 been proposed that this enzyme may interact directly or indirectly with the CesA complex, thereby facilitating the channelling of UDP-Glc to the catalytic CesA machinery (Amor et al., 1995). In addition, the overexpression of the cotton SuSy gene in poplar lead to an increase in cellulose formation (Coleman et al., 2009), suggesting that SuSy is involved in the production of UDP-glucose for CesA.

It has not been firmly demonstrated that a primer is required to initiate cellulose polymerization. The role of cellobiose as a primer has been investigated, but results suggest that the disaccharide acts as an activator rather than a primer (Li and Brown 1993; Lai Kee Him et al., 2001). Sitosterol-β-glucoside was also shown to be used as an acceptor by a plant CesA in an in vitro experiment (Peng et al., 2002). However, these findings have yet to be confirmed in vivo. Further knock-out mutants of the Arabidopsis sitosterol pathway do not show any deficiency in cellulose, thus, the role of sitosterol-β-glucoside as a primer remains controversial (Somerville’s group, personal communication). More research in this area is needed as Shrick et al. (2004) reported contrasting results, where a decrease of cellulose in the walls of Arabidopsis sterol biosynthesis mutants was found.

While the requirement of a primer for cellulose biosynthesis remains unclear, there are strong experimental evidence in bacteria and plants that cellulose chain elongation occurs from the non-reducing end of the molecule (Koyama et al., 1997; Lai Kee Him et al., 2002). The same observation has been made for chitin biosynthesis in diatoms (Imai et al., 2003). The elongation of cellulose chains is most likely initiated in the cytoplasm as sucrose synthase and UDP-Glc pyrophosphorylase both occur in this compartment. This is further supported by the recent structural data obtained on the Rhodobacter CesA showing the existence of a channel in the enzyme catalytic subunit that is able to translocate a growing chain from the cytoplasm to the periplasmic space (Morgan et al., 2013). Prior to the publication of the work on Rhodobacter, a number of models for the extrusion of cellulose chains across the plant plasma membrane were proposed. These models involving porin- like proteins, flippases (Brown and Saxena, 2000) or the formation of a pore by the transmembrane domains of each CesA (Delmer 1999) are likely incorrect. Instead, analysis of the bacterial and predicted plant CesA structures (Sethaphong et al.,

34 2013) support the existence of a comparable passage in both types of enzymes for the extrusion of cellulose chains.

It appears the assembly of cellulose chains into microfibrils is a spontaneous process as no proteins have been linked to this step of cellulose formation (Guerriero et al., 2010). Microfibrils are composed of both well ordered crystalline and less ordered non-crystalline regions. Several techniques (transmission electron microscopy, X-ray diffraction and solid-state NMR spectroscopy) have shown that the width of cellulose microfibrils in plant primary cell walls is in the range of 2-4 nm (Roland et al. 1975; Chanzy et al. 1978, 1979; Revol et al. 1987; Emons 1988; Boylston and Hebert 1995; Ha et al. 1998; Thimm et al. 2002; Kennedy et al. 2007). Detergent extracts of the plasma membranes of blackberry cells have also been found to produce microfibrils in vitro of similar width (Lai Kee Him et al., 2002). Interestingly, the microfibrils synthesized in vitro were 10% more crystalline than their counterpart in primary cell walls (40 vs 30%) (Lai Kee Him et al., 2002). This may be due to the absence of other carbohydrates in the in vitro system, favouring the perfect alignment and packing of chains in microfibrils. The degree of cellulose crystallinity in the oomycete S. monoica was reported to be slightly poorer (≤ 25%) than in the plant system, as measured by X-ray diffraction (Bulone et al., 1992).

Cellulose biosynthesis probably involves proteins other than CesAs. A number of plant proteins (e.g. KORRIGAN, KOBITO1 and COBRA) have indeed been proposed to play a role in cellulose biosynthesis through the analysis of mutants in Arabidopsis. In most cases, the function of these proteins in cellulose biosynthesis remains poorly understood and lacks direct proof-of-function. As an example, the gene korrigan (Kor) encodes a putative membrane-bound β-(1,4)-endoglucanase and Kor mutants show a decrease in cellulose by 40% compared to the wild-type plants (Sato et al., 2001). Despite the many studies on different KORRIGAN genes from various species and the observation of different phenotypes in Kor mutants, the precise function of the protein remains elusive.

35 1.7.3 Cellulose synthase genes from oomycetes

Cellulose synthases are processive GTs responsible for the formation of cellulose chains. As mentioned earlier, they belong to the GT2 family.

The sequencing of the P. infestans genome facilitated the identification of oomycete CesAs (Tyler et al., 2006b; Haas et al., 2009). Four CesA genes designated PiCesA1-4 were functionally characterized in P. infestans (Grenville-Briggs et al., 2008). Soon after, the three S. monoica orthologs of the P. infestans CesA genes, were identified (SmCesA2, SmCesA3 and SmCesA4) (Fugelstad et al., 2009). Interestingly, no equivalent P. infestans CesA1 was found in S. monoica. Recently, six CesA genes were found in the genome of S. parasitica (Jiang et al, 2013).

The function of the P. infestans CesA genes was investigated by RT-PCR at different developmental stages of the microorganism, via gene silencing approach and treatment with 2,6-dichlorobenzonitrile (DCB) (Grenville-Briggs et al., 2008). All CesA genes were shown to be upregulated during cyst germination and the formation of appressoria. Silencing of the four CesA genes by RNAi resulted in changes of the cell wall structure. As a consequence, P. infestans was no longer able to successfully infect the host plant. The authors showed that the formation of a normal appressorium is required for pathogenesis and infection was not possible without functional CesA genes. The importance of cellulose in P. infestans infection was further supported by the use of DCB, an inhibitor of cellulose biosynthesis, showing evidence of cellulose in facilitating infection of the host plant. Cysts can germinate in the presence of DCB, but the abberrant appressoria-like structures which form in the presence of the drug are unable to penetrate the host. It is noteworthy that all experiments in this study analyzed the combined silencing of all four CesAs simultaneously. Therefore, the function of each CesA gene remains to be determined by analyzing them individually. Mycelium growth in S. monoica was decreased when cells were cultivated in the presence of DCB (Fugelstad et al., 2009). RT-PCR analysis of the treated and control cells showed the mycelium reacts to DCB by increasing the expression of all its SmCesA genes (Fugelstad et al., 2009). These data, together with a higher β-glucan synthase activity in vitro, suggest that the cells

36 can adapt to the presence of DCB through a compensatory mechanism (Fugelstad et al., 2009).

Sequence analysis (Fugelstad et al., 2009) and domain comparison (Figure 15) of CesA genes from different oomycetes show the presence of the D,D,D, QXXRW motif found in most processive GTs (Campbell et al., 1997). Moreover, Figure 15 shows the presence of a domain specific to the oomycetes called pleckstrin homology (PH) domain (Grenville-Briggs et al., 2008; Fugelstad et al., 2009). This domain is located at the N-terminal end of some of the oomycete CesAs (Figure 15). SmCesA2 in S. monoica and PiCesA1, PiCesA2 and PiCesA4 in P. infestans all contain PH domains (Grenville-Briggs et al., 2008; Fugelstad et al., 2009). Although the full length sequence of SmCesA4 is not available, a PH domain is expected to occur at the N-terminal of the gene product as for the PiCesA4 ortholog. None of the CesA3 harbor this type of domain (Figure 15). Instead, SmCesA3 and its orthologs PiCesA3 and SpCesA3 contain additional transmembrane domains (Grenville-Briggs et al., 2008; Fugelstad et al., 2009). It is noteworthy that the occurrence of PH domains has never been reported in any other carbohydrate synthase, including CesAs from plants and bacteria (Figure 15A). The function of the PH domain will be discussed in section 1.11.

Figure 15. Domain organisation of CesA from a diverse range of organisms (diagram constructed using DomainDraw (Fink and Hamilton, 2007)). A. Domain comparison between oomycete, plant and bacterial CesAs shows the presence of PH (blue) and zinc finger (black) domains unique to the oomycetes and plants, respectively (Fugelstad et al., 2009). B. Domain organisation of the S. parasitica CesAs shows the occurrence of a PH domain at the N-terminal of the same ortholog proteins from other oomycete species (Figure 15A) and the absence of a PH domain in CesA3 (Díaz- Moreno et al., unpublished results).

38 1.8 Overview of the secretory and endocytosis pathways of proteins

As earlier discussed in section 1.6.2 and 1.6.4.3, a distinct MIT domain is found in Chs. Based on our research, the MIT domains studied in this thesis might be involved in the trafficking of chitin synthases and details of this discussion will be further described in the results section 3.3. Therefore, in this section, a brief overview of the secretory and endocytosis pathways is presented.

1.8.1 Secretory pathway

Cells are constantly producing, degrading and recycling proteins to respond to the metabolic and structural needs of the organism. These processes involve specific intracellular trafficking that is dependent on clathrin-coated vesicles (CCVs). In the case of transmembrane proteins, the secretion by clathrin-coated vesicles begins by an interaction of the proteins with adaptor protein (AP) complexes via specific sorting signals (see section 1.8.2). The sorting signals (“dileucine-based” and “tyrosine- based” motifs) present at the cytosolic tail of the transmembrane proteins are recognized by a specific subunit of the AP (Sandoval and Bakke, 1994; Mellman, 1996). While the “dileucine-based” motif is part of the signature peptide [D/E]XXXL[L/I]1 involved in protein targetting (Bonifacino and Traub, 2003), for the “tyrosine-based” motifs, there are two types: the YXXΦ2 and the NPXY3 motifs (Bonifacino and Traub, 2003). The YXXΦ motif, which binds to all APs, but with variable affinities (Owen et al., 2004) occurs in many transmembrane proteins targeted to endosomes and lysosomes (Bonifacino and Traub, 2003) (Figure 17). Once a specific sorting signal is recognized, proteins are transported by three main types of coated vesicles which can be distinguished by their coat type: clathrin, coat protein I (COPI) and coat protein II (COPII). Only the clathrin coat is discussed in this thesis.

1 D/E corresponds to aspartic and glutamic acids, X represents any amino acid residue and L/I correspond to leucine and isoleucine respectively 2 Y: tyrosine, X: any amino acid, Φ: amino acid with a hydrophobic side chain such as phenylalanine, leucine, isoleucine, methionine and valine 3 N: asparagine, P: proline

39 CCVs form through the assembly of triskelions, which consist of clathrin heavy and light chains and form a polyhedral outer scaffold (Figure 16A). The inner layer of CCVs consists of AP complexes that interact with the target protein and the phospholipid bilayer. The triskelion scaffold forms a stable structure that protects the vesicles transporting the cargo proteins. CCVs are involved in the secretory pathway, which, transports proteins from the trans-Golgi to their final destination. During trafficking, proteins are sorted and addressed to their target cell compartment. In addition to this forward trafficking, some proteins are transported back from the Golgi apparatus to the ER, following the so-called retrograde process. This typically applies to proteins that are resident of the ER but whose function requires modifications that take place in the Golgi apparatus (e.g. N-glycan processing).

Secreted proteins traffic through the secretion pathway and are delivered to the extracellular environment by exocytosis following membrane fusion between the transport vesicles and the plasma membrane (Rohn et al., 2000). In addition to contributing to exocytosis, CCVs are part of the endocytosis pathway (Figure 16).

Figure 16. Endocytosis process. A. Principal steps involved in the formation of clathrin-coated vesicles (CCVs). Adaptor protein 2 (AP2) recognizes a specific sorting signal on a cargo protein via its µ2 subunit. Once attached to it, the three heavy chains (CHC) and three light clathrin chains (CLC) assemble by interacting together and with the ears of the AP. The release of the CCV is performed by dynamin, a GTPase that allows membrane fission. B. Depolymerization of CCVs and targeting of their cargo proteins. The uncoating process allows the endocytic vesicles to fuse to the early endosomes (sorting endosomes). At this point, proteins can be recycled back to the PM or internalized inside the endosome to form multivesicular bodies (MVBs), a process that involves the ESCRT machinery. Further sorting occurs in the MVBs where proteins can be targeted to the Golgi apparatus or the lysosome where they are degraded. (adapted from Tauber, 2003 and Winter and Hauser, 2006).

41 1.8.2 Clathrin-mediated endocytosis pathway

The endocytosis pathway may occur in a clathrin dependent or independent manner. In the case of the CCV-mediated endocytosis, the process is triggered by cell signals that initiate the assembly of the coat. Proteins that are recycled or degraded contain a cytosolic sorting signal (described in section 1.8.1) recognized by a specific subunit of the membrane-bound AP complexes.

In addition to the well-identified AP complexes in mammalian (AP1, AP2, AP3, AP4) and yeast (AP1, AP2, AP3) (Robinson and Bonifacino, 2001; Owen et al., 2004), a new AP (AP5) was recently described (Hirst et al., 2011). It occurs in most eukaryotic organisms with the exception of fungi and stramenopiles (Hirst et al., 2011).

Figure 17. Representation of the adaptor protein complexes and their subunits. Specific subunits are known to interact with sorting motifs from cargo proteins and with clathtrin. For instance, in AP3 the subunit complex δ/σ or σ itself interacts with dileucine-based motifs ([D/E]xxxL[L/I]) and the µ subunit interacts with tyrosine-based motifs (YxxΦ) in the cargo protein. Note that these two sorting motifs can also interact with other adaptor proteins (adapted from Braulke and Bonifacino 2009).

Each form of AP complex is involved in a different type of vesicle trafficking. For the purpose of this thesis only AP2 and AP3 are discussed (Figure 17). AP2 acts at the plasma membrane level and mediates endocytosis (Owen et al., 2004; Jackson et al., 2010). AP3 transports proteins from the endosomes to the lysosomes. In mammalian cells, like fibroblasts, AP3 is responsible for targeting lysosomal-

42 associated proteins to the lysosomes (Dell’Angelica et al., 1999; Le Borgne et al., 1998) and in yeast, for transporting proteins from the TGN to the vacuoles (Cowles et al., 1997; Odorizzi et al., 1998). A minor fraction of AP3 was also localized at the trans-Golgi (Peden et al., 2004). As opposed to AP1 and AP2 that clearly associate with clathrin, the in vivo and in vitro results obtained on AP3 are conflicting (Newell- Litwa et al., 2007).

In AP2-mediated endocytosis, the sorting signal of the target protein is first recognized by AP2 at the plasma membrane (Figure 16A). The vesicle is then stabilized through interactions between the globular domain of the clathrin heavy chain and a specific subunit of AP2 (Pearse and Robinson, 1990; Bonifacino and Traub, 2003). The CCV buds from the donor membrane thanks to the action of dynamin, which wraps around the bud of the vesicle. The energy released by dynamin via the hydrolysis of GTP allows the vesicle to separate from the membrane (Takel et al., 1995). The vesicle subsequently moves toward its final destination. The clathrin coat must be depolymerized for the vesicle to fuse with the acceptor membrane (Figure 16B). This process is performed by HSc70, i.e. a chaperon protein that allows the recycling of the triskelions (Eisenberg and Greene, 2007). Vesicles fuse to early endosomes, forming tubulovesicular structures that act as sorting platforms (William and Urbé, 2007). At this stage some proteins can be recycled and returned to the PM or they are internalized inside the endosome via the Endosomal Sorting Complexes Required for Transport (ESCRT). Once in the multivesicular bodies, proteins are either recycled or transported to lysosomes.

1.9 Microtubule interacting and trafficking (MIT) domains

The present investigation involved the characterization of the MIT domain present in both chitin synthases (Sm_MIT1 and Sm_MIT2) from S. monoica. Sections 1.9 and 1.10 describe the properties of MIT domains, its interaction with the MIT-interacting domains (MIM) and examples of proteins containing MIT domains, including chitin synthases.

43 The terms “Microtubule Interacting and Trafficking domain” and “Microtubule Interacting and Transport domain” both abbreviated as (MIT) were introduced to describe protein domains involved in microtubule binding and intracellular transport of proteins (Ciccarelli et al., 2003; Alonso, 2011). Both terms refer to the same domains and are therefore interchangeable.

MIT domains have an average mass slightly lower than 10 kDa and their sequences are often poorly conserved, which makes their identification by sequence similarity difficult (Rigden, 2009). Prediction of their binding specificity through bioinformatic approaches has also proven to be challenging (Hurley, 2011). The occurrence of conserved key structural residues led to the hypothesis that MIT domains share a similar 3D structure (Scott et al., 2005b). The 3D structure of the human AAA ATPases VPS4A and VPS4B revealed 3 asymetric α-helices linked by short loops (Scott et al., 2005a; Scott et al., 2005b; Takasu et al., 2005) (Figure 18).

Figure 18. Structure of the MIT domain from the human VPS4A protein (from Scott et al., 2005a).

Several reports have shown that MIT domains bind amphipathic helices named MIT- interacting motifs (MIMs) (Stuchell-Brereton et al., 2007; Obita et al., 2007; Hurley and Yang, 2008; Lee et al., 2012). These low molecular weight proteins exhibit a basic N-terminus and an acidic C-terminus that interact together (“basic-acid terminus”). MIMs are typically present in a family of structurally related proteins called charged multivesicular body proteins (CHMPs) and CHMPs are components of the ESCRT-III complex (William and Urbé, 2007). For instance, MIM motifs allow the

44 recruitment of MIT domains proteins such as Vps4, which, are needed at the ESCRT- III complex to catalyse the disassembly of the complex.

Few MIT-containing proteins have been fully characterized and our knowledge is mostly based on the human and yeast examples that will be described in section 1.10. MIT domains typically occur in proteins involved in cellular trafficking, where they act as protein-interacting modules (Stuchell-Brereton et al., 2007). In most cases, MIT-containing proteins are enzymes with AAA ATPase domains. AAA ATPase domains are involved in different types of cellular functions (Vale, 2000). For instance, the human VPS4 binds the ESCRT-III complex, whereas spastin binds to microtubules. Some other MIT containing enzymes are devoid of AAA ATPase domains, as is the case for the de-ubiquitinating enzymes AMSH and USP8 (also known as UBPY) (Komander et al., 2009; Hurley and Stenmark, 2011). Some enzymes like calpain 7 (protease) and Vta1 contain two MIT domains and interact with ESCRT-III. More details are provided in section 1.10 regarding the role of MIT domains in carbohydrate-active enzyme and other MIT-containing proteins.

1.10 MIT domains in carbohydrate-active enzymes and other types of proteins

In this section, different types of enzymes that contain MIT domains are presented to illustrate the range of functions in which MIT domains can be involved.

1.10.1 Carbohydrate-active enzymes

Both Chs from S. monoica were shown to each contain a single MIT domain. The function of these domains in carbohydrate-active enzymes is unknown (Guerriero et al., 2010). More recently, it was shown that two out of the six Chs of S. parasitica also contain N-terminal putative MIT domains (Rzeszutek et al., 2015, manuscript) and only one of the two putative Chs genes from Aphanomyces euteiches contains a putative MIT domain. The presence of MIT domains in oomycete Chs but not in yeast and fungal enzymes makes these domains interesting to study as they may have a specific function in oomycetes which is perhaps performed by separate proteins in

45 other eukaryotic micro-organisms. The sequence alignment of MIT domains from carbohydrate active enzymes (Chs) and non-carbohydrate active enzymes (VPS4) reveals several conserved amino acid residues (Figure 19). The alignment also highlights the presence of a dileucine motif in S. parasitica and S. monoica Chs2 suggesting the proteins can bind to AP2 and are internalized during the endocytosis process. Interestingly, there is a proline residue at the -1 position from the first leucine. Its presence has been found to increase the binding with AP3 (Rodionov et al., 2002).

Figure 19. Sequence alignment of MIT domains from oomycete Chs and other proteins. The MIT domain consensus sequence as described by Scott et al. (2005) is shown in red bold letters. Dileucine-based motifs are shown in blue. The predicted secondary structure of MIT domains is shown above the sequences (helices H1-3) based on the NMR structure of Sm_MIT1 (Brown et al., 2015).

1.10.2 Non-carbohydrate-active enzymes

1.10.2.1 Spastin

Spastin is an AAA+ family member that causes spastic paraplegia in humans (HSP) (Taylor, 2012). The AAA domain sequence of spastin is closely related to the AAA domain of kanatin (p60), which is a microtubule severing enzyme. However, spastin and kanatin have no significant sequence similarity in their N-terminal regions (Roll- Mecak et al., 2008). Early studies suggested a similar function for the two proteins as they are both able to bind microtubules via their N-terminal regions (Errico et al., 2002). Recently, the microtubule severing function of spastin was demonstrated (Evans et al., 2005; Roll-Mecak et al., 2008). However, the MIT domain of spastin does not seem to be involved in microtubule binding. Indeed, studies on truncated

46 spastin show that the N-terminal region of the protein contains a microtubule-binding domain (MTBD) different from the MIT that is responsible for the actual binding of the protein to microtubules and its microtubule severing activity (White et al., 2007). The MTBD domain binds tubulin in an ATP-independent manner (White et al., 2007) and the disruption of the microtubule structure occurs by ATP hydrolysis that is catalysed by the C-terminal AAA ATPase domain of spastin (Evans, 2005). The MIT domain of the protein is likely responsible for targeting spastin at its site of action as mutations in the MIT domain of spastin prevent its recruitment to the midbody during cytokinesis (Connell et al., 2009). As a consequence, microtubules remain intact affecting cell division of the two daughter cells.

1.10.2.2 De-ubiquitinating enzymes

The MIT domain of de-ubiquitinating enzymes, AMSH (Tsang et al., 2006) and USP8 (Row et al., 2007) allows the proteins to be targeted to the ESCRT-III complex (Hurley and Stenmark, 2011). Also noteworthy, the USP8 MIT domain from mouse was found to bind microtubules in vitro and also co-localized with microtubules in vivo suggesting the potential role of microtubules in assisting the targeting of the de- ubiquitinating enzymes (Berruti, 2010).

1.10.2.3 VPS4 (Vps4)

The vacuolar protein sorting Vps4 has been studied extensively due to its key role in multivesicular bodies biosynthesis and in other processes like cytokinesis and viral budding where ESCRT-III is involved (Stuchell-Brereton et al., 2007). Vps4 is a member of the AAA+ ATPase family and there is a single isoform of the Vps4 protein in yeast, whereas, two isoforms named VPS4A and VPS4B occur in humans. Structural studies on Vps4 from both organisms showed that the MIT domain binds to specific proteins of the ESCRT-III complexes containing the MIT-interacting motifs. These interactions allow Vps4 to be recruited during MVB formation. The energy produced by ATP hydrolysis allows the ESCRT-III complex to dissociate from the endosomal membrane (Babst et al., 2002) and the components of the complex are then recycled for a next round of complex assembly (Hurley and Hanson, 2010).

47 1.11 Pleckstrin homology domains

In oomycetes, a distinctive domain is also found in the CesA protein, called pleckstrin homology domain (section 1.7.3). This thesis presents the functional characterization of the pleckstrin homology domain found in S. monoica CesA2. Described in the sections below are some background on PH domain and the membrane lipid bilayer.

The membrane lipid bilayer delimiting the cell is composed of lipids, essentially phosphatidylethanolamine (PE), phosphatidylcholine (PC), phosphatidylserine (PS), cholesterol and sphingomyelin. Phosphoinositides (PtdIns) and phosphatidic acid (PA) are present in smaller amounts in the cell membrane and are more specifically involved in signalling. Some cellular lipids are zwitterions at pH 7.0, e.g. PC and PE, while others are negatively charged, e.g. PS, PA and PtdIns, thereby creating a negative electrostatic potential which facilitates the recruitment of proteins by interacting with specific protein modules such as the PH domains. Pleckstrin homology domains (PH) represent the most important family of phosphoinositide- binding modules (Lemmon 2008).

1.11.1 Background

In S. cerevisiae, there are 30 known proteins that contain PH domains, compared to 696 proteins in humans (SMART database, 2014). The PH domain was first discovered in a protein called pleckstrin, which is the major substrate of protein kinase C in platelets (Tyers et al., 1988). Although, most PH-containing proteins have only one such PH domain, pleckstrin contains two, one at its N-terminal and the second at its C-terminal end (Tyers et al., 1988). PH domains occur in proteins with diverse functions, for instance in proteins involved in membrane trafficking, phospholipid modification, cytoskeletal organization and signal transduction (Ingley et al., 1994; Musacchio et al., 1993).

48 1.11.2 Structure of PH domains

PH domains typically consist of around 120 amino acids. Sequence identity between different PH domains is in the range 7-23% only. However, their 3D structure is highly conserved (Lemmon and Ferguson, 2000). The first PH domain structures were obtained from the mouse brain β-spectrin (Macias et al., 1994) and pleckstrin protein (Yoon et al., 1994). To date, there are more than 100 structures of PH domains available publically in Protein Data Bank database (Kutateladze, 2010). Figure 20 shows that the typical structure of PH domains consists of 7 antiparallel β-strands and one α-helix at the C-terminal which blocks the structure on one side (Lemmon and Ferguson 2000).

Figure 20. Structure of the C-terminal PH domain of human pleckstrin (PDB 1XX0) (adapted from Edlich et al., 2005).

The β-strands are connected by loops of variable length and sequence. The structure of the PH domain from β-spectrin was resolved by NMR and revealed that the molecule is electrostatically polarized and between the two polarized faces contain a pocket that could serve for ligand binding (Macias et al., 1994) (Figure 21). Similar polarity patterns occur in other PH domains as is the case in the PH domain of dynamin (Ferguson et al., 1994). The conserved region that includes the C-terminal α-helix is charged negatively, whereas the variable loops are exposed on the positively charged face. The length and sequence variability of the loops connecting the β-strands are possibly at the origin of the observed range of affinities and specificities between different PH domains (Ferguson et al., 1994). However, none of these loops seem to affect the overall fold of the domain (Yoon et al., 1994). Analysis of the structure of the N-terminal PH domain of pleckstrin demonstrated the presence of a key Trp (92) in the C-terminal α-helix, which was later shown to be conserved in

49 all PH domains. It has been suggested that this amino acid plays a role in stabilizing the PH domain by interacting with the hydrophobic core β-barrel (Yoon et al., 1994). Moreover, 4 of the 6 lysine residues that occur at the β-barrel entry are conserved in many proteins containing PH domains (Haslam et al., 1993; Mayer et al., 1993; Musacchio et al., 1993; Lemmon, 2008).

Figure 21. Electrostatic potential of β-spectrin. The red color corresponds to the negatively charged face, whereas the purple color highlights the positively charged face. The variable loops in white are mostly part of the positively charged face and are thought to form the binding surface of the PH domain (adapted from Macias et al., 1994).

1.11.3 Phosphatidylinositol

Lipid vesicles containing phosphatidylinositol biphosphate were used to demonstrate pleckstrin interactions with PtdIns through the PH domain (Harlan et al, 1994). PtdIns are phosphorylated derivatives of phophatidylinositol and occur in several forms, i.e. phosphatidylinositol monophosphate (PIP), phosphatidylinositol biphosphate (PIP2) and phosphatidylinositol triphosphate (PIP3). A higher number of phosphate groups on the PtdIns molecules increases the strength of the binding to the PH domain (Takeuchi et al., 1997). PtdIns consist of a glycerol backbone esterified by two fatty acids and a phosphate group. The latter connects the non-polar part of the molecule to a myo-inositol group via a phosphodiester bond (Figure 22) making the molecule amphiphatic. The inositol group can be further phosphorylated by different types of kinases, leading to a total of seven derivatives of phosphatidylinositol.

Figure 22. Structure of a phosphatidylinositol.

The levels and turnover of PtdIns are tightly regulated during cell growth. The different types of PtdIns are preferentially distributed in different membrane compartments (Figure 23) (Le Roy and Wrana, 2005; Kutateladze, 2010). Signal transduction processes can be finely tuned through the control of the turnover of phospholipids in the membrane by kinases, phosphatases and phospholipases. In mammalian cells, PtdIns represent a small proportion of the total phospholipids (Lemmon 2008). However, they play important cellular roles, for instance in the interaction between phospholipid-binding proteins and the cell membrane (Di Paolo and De Camilli, 2006).

Figure 23. Specific distribution of phosphatidylinositides (PtdIns) in an animal cell. PtdIns allow the recruitment of specific proteins to distinct intracellular compartments. EE, early endosomes, MVB multivesicular bodies (adapted from Kutateladze et al., 2010).

1.11.4 Function and mode of interaction of PH domains

While most of the PH domain-containing proteins interact with the plasma membrane at some stage to perform their function, the precise role of many PH domains remains unclear (Lemmon and Ferguson, 2001). The studies available suggest that PH domains are involved in targeting their corresponding proteins via either protein- protein or protein-lipid interactions. These interactions take place at the cytosolic face of the membrane (Mayer et al., 1993; Lemmon and Ferguson, 2000; DiNitto and Lambright, 2006). Both protein-protein and protein-lipid interactions can be established simultaneously if different interaction surface is present as is the case for the PH domain of the G protein-coupled receptor kinase GRK2 (Lodowski et al., 2003). While these interactions are well known to occur at the membrane, the mechanisms that allow the host proteins to reach the lipid bilayer remain poorly understood. Apart from a few exceptions, in vivo studies in S. cerevisiae have generally failed to demonstrate that the PH domain itself can target the protein to the membrane (Yu et al., 2004).

52 Most PH domains seem to interact with PtdIns essentially through nonspecific interactions. As a consequence, they are able to bind to more than one type of phospholipid. Nonspecific interactions could involve charges, amphiphilicity and membrane curvature. (Lemmon 2008). This lack of specificity allows the proteins that contain nonspecific PH domains to occur in different intracellular membranes. PH domains that interact with PtdIns through weak interactions may oligomerize to enhance their binding to the membrane or interact with a second ligand that assists binding to PtdIns (Carlton and Cullen, 2005). The participation of two different ligands would restrict the targeting of the protein and the site of its activity, as both ligands would have to be present at the same time to allow protein attachment.

Of the mammalian PH domains characterized to-date, only 10% exhibit a high affinity and specificity for PtdIns (Lemmon and Ferguson, 2000). In S. cerevisiae, this percentage is even lower as only one out of 33 PH domains investigated binds PtdIns with high affinity and specificity. Most of the remaining PH domains interact with PtdIns in a promiscuous manner and a small group do not show any binding to PtdIns at all (Yu et al., 2004).

A common feature of PH domains that do show high affinity interactions for

PtdIns(3,4)P2 and PtdIns(3,4,5)P3 is in the presence of the (KXn(K/R)XR) motif, which occurs in the variable loop between the β1 and β2 strands (Isakoff et al., 1998; Dowler et al., 2000; Lemmon 2008). The basic amino acids lysine and arginine are responsible for most of the interaction with the phosphate group of the PtdIns. Both in vitro binding to PtdIns and cell membrane targeting were affected by mutations in the key basic residues of the (KXn(K/R)XR) motif (Lemmon and Ferguson 2001), suggesting a key role for these amino acids in the binding. As is the case for other membrane-binding domains, PH domains contain a pocket defined by the variable loops that is responsible for the stereospecific recognition of unique phospholipid species and their soluble headgroups (Hurley, 2006). Although other loops can be involved in the specific interactions, the β1-β2 loops are usually responsible for the establishment of the contact with the PtdIns headgroups.

The first well characterized high-specificity PH domain is from phospholipase C

(PLC) δ1. It contains a stereospecific recognition site for PtdIns(4,5)P2 (Lemmon et

53 al., 1995; Garcia et al., 1995; Kavran et al.,1998). The Bruton's tyrosine kinase (Btk) PH domain is also highly specific but for PtdIns(3,4,5)P3 (Rameh et al., 1997a; Kojima et al., 1997; Várnai et al., 1999). High-specificity PH domains fused with green fluorescent protein (GFP) can be used to identify the intracellular localization and determine the levels of PtdIns in the cell (Balla et al., 2000; Halet, 2005) (Figure 24). The stereospecificity of the PH domain-PtdIns interactions may be increased in some cases by the electrostatic potential (nonspecific interactions). Conversely, in some cases a change of electrostatic potential may decrease binding to PtdIns (Hurley and Misra, 2000; DiNitto et al., 2003). It has been proposed that nonspecific interactions assist the overall interaction by adequately orienting the domain with respect to the membrane surface (DiNitto et al., 2003). Hydrophobic side chains in the PH domain contribute to an increased affinity with the membrane by facilitating the penetration of the domain in the lipid bilayer (Hurley and Misra, 2000).

Figure 24. Tools to vizualize and localize PtdIns in cell membranes. PH domains with high affinity and specificity fused to GFP can be used to localize the different types of PtdIns. Protein abbreviations: four-phosphate-adaptor protein (FAPP1), oxysterol-binding protein (OSBP), tandem PH domain- containing protein (TAPP1), protein kinase B (PKB), general receptor for phosphoinositide 1 (GRP1), Bruton’s tyrosine kinase (Btk), phospholipase C (PLC δ1) (adapted from Kutateladze 2010).

54 1.11.5 Classification of PH domains

Table II shows a new and simple classification of the different types of PH domains (Maffucci and Falasca, 2001), which was proposed to replace the one previously published by Rameh et al. (1997). The new classification is based on 3 different groups of PH domains, which can be distinguished by their different affinities with respect to their ligands. Briefly, group 1 consists of PH domains that interact strongly and specifically with PtdIns, group 2 includes domains with weak and promiscuous interactions with PtdIns and group 3 contains domains that establish non-specific interactions with PtdIns.

Table 2. Classification of PH domains as proposed by Maffucci et al. (2001). PH domains are classified based on their affinity and selectivity to PtdIns. Protein abbreviations: Bruton’s tyrosine kinase (Btk), phospholipase C δ1 (PLC-δ1), general receptor for phosphoinositide 1 (Grp1), phospholipase C β1 (PLC-β1), diffuse B-cell lymphoma (Dbl), GRB2-associated-binding protein 1 (Gab1), insulin receptor substrate 1 (IRS-1), phospholipase C γ1 (PLC-γ1), diacylglycerol kinase-N (DAG K-δ).

55 2 Aims of the present investigation ______

The main objective of this thesis was to characterize the cellulose synthase pleckstrin homology (PH) domain and the chitin synthase microtubule interacting and trafficking (MIT) domain that are involved in cell wall formation of oomycete Saprolegnia monoica. To the best of my knowledge, the domains described herein have never been reported in any known glycosyltransferase. In addition, there is to-date no information on their functions in oomycetes in relation to cell wall formation, although putative functions have been assigned to animal orthologs for these domains. Deciphering the functions of cellulose synthase pleckstrin homology domain and the chitin synthase microtubule interacting and trafficking domain in pathogenic oomycetes may facilitate the development of new methods for disease control.

2.1 The specific aims of the thesis were:

• The implementation of in vitro assays to study membrane-bound glycosyltransferase activities involved in cell wall biosynthesis (Paper I).

• The functional characterization of the pleckstrin homology (PH) domain of the cellulose synthase 2 from S. monoica (SmCesA2) (Paper II).

• The expression, purification and structural characterization of the chitin synthase 1 and 2 MIT domains (SmChs1 and SmChs2) from S. monoica (Paper III).

• The modelling of the interactions between lipids and the microtubule interacting and trafficking (MIT) domain of the chitin synthase 1 (SmChs1) from S. monoica (Paper IV).

• The characterization of two putative chitin synthases (AeChs1 and AeChs2) from microsomal fractions of Aphanomyces euteiches (Paper V).

56 3 Results and discussion ______

3.1 Glycosyltransferase assays (Paper I)

A key function of numerous glycosyltransferases (GTs) from eukaryotic microorganisms is the biosynthesis of cell wall polysaccharides which are polymerized through the transfer of monosaccharides to elongating glycan chains. Understanding the cell wall formation by GTs in pathogenic microbial species can provide important information for disease control. Indeed, this knowledge can be exploited to design inhibitors against GTs vital for the survival of pathogenic fungi, oomycetes and bacteria, which infect humans, animals and plants. An important tool for the characterization of GTs is the development of specific in vitro assays. Relevant biochemical procedures are described in detail in paper I. Most GTs have not been biochemically characterized, mostly because of their instability after extraction from their membrane environment and the difficulty of expressing them recombinantly in a catalytically active form. Even when recombinant expression is successful, finding the optimal conditions to perform the experiments remains a challenge as the requirements for sugar donors, acceptors and effectors are largely unknown. Furthermore, many enzymes form complexes and the identity of their partners, which may be required for the enzymes studied to be active, is often unknown. Despite these difficulties, a number of GTs have been functionally characterized. Examples include chitin synthases from S. cerevisiae (Martínez- Rucobo et al., 2009) and S. monoica (Guerriero et al., 2010). These enzymes were expressed in bacteria and yeast, respectively, and purified after detergent extraction from the plasma membrane. Structural analysis of the polymer products formed in vitro confirmed the type of reactions catalyzed by the recombinant enzymes.

Paper I describes two types of in vitro assays for GTs, one based on spectrophotometric approach and the other a radiometric approach. Both assays can be used for processive and nonprocessive GTs. The examples provided correspond to plant GTs, but the same approach can be readily applied to any microbial enzyme

57 by using the appropriate sugar donors and acceptors. This is illustrated in paper V where the radiometric assay was applied to oomycetes chitin synthases.

Although less sensitive, the spectrophotometric assay has some advantages over the radiometric assay. The most obvious is the non-requirement of radio-isotopes which must be used under specific laboratory environments and involves a more stringent waste management. However, in order for the spectrophotometric assay to be a viable option, the enzyme tested must exhibit a high activity and needs to be highly enriched or purified. In this approach, the spectrophotometric assay involved a combination of three different enzymes, i.e. the GT of interest, pyruvate kinase (PK) and lactate dehydrogenase (LDH) (Figure 2, paper I). These enzymes are typically sensitive to impurities, and crude extracts from the plasma membrane may contain interfering activities such as oxidoreductases and phosphatases, which might affect the LDH and PK reactions.

The radiometric assay (Figure 25) is less demanding in terms of enzyme purity and allows the study of GTs in crude protein preparations such as microsomal fractions or detergent extracts of membrane fractions. In this type of assay, the radiolabeled monosaccharide from the substrate (NDP-sugar in which the monosaccharide is labelled with either 14C or 3H) is detected by scintillation counting after the radiolabelled ethanol-insoluble polymers (product) were retained on glass-fiber filters (Figure 25A).

58 A B

Figure 25. Radiometric assays. A. Workflow to assay for GTs that form ethanol-insoluble product. B. Workflow to assay for GTs that form soluble product.

For enzymes that form ethanol-soluble glucans, the nucleotide-sugar used as a substrate and the enzyme product must be separated before measurement can be conducted for the radioactivity incorporated into the newly synthesized glycan. This purification step is performed by using anion-exchange chromatography during which the excess of NDP-sugar and the NDP released by the enzyme as a secondary product are retained on the column, while the soluble radioactive glycan formed by the GT is readily eluted. The eluate can then be analysed by liquid scintillation counting (Figure 25B). Importantly, this method can be used only for GTs that form neutral saccharides, which is the case of most enzymes, with the exception of GTs that catalyze the transfer of uronic acids to glycan chains. In all cases, a series of adapted controls must be performed as detailed in Table 2 of paper I. Furthermore, product characterization must be performed to confirm the type of reaction catalyzed by the GT, i.e. the type of glycosidic bond formed. This can be achieved using enzymatic or chemical methods, as explained in Bulone et al. (1995). Given the availability of highly specific hydrolytic enzymes, a simple reaction based on glycoside hydrolases is performed. More advanced techniques including linkage analysis by permethylation, 13C-NMR spectroscopy, mass spectrometry and X-ray diffraction can also be used depending on the type and quantity of glycan synthesized.

59 3.2 Characterization of the PH domain from SmCesA2 (Paper II)

PH domains have been studied primarily in mammalians but are also found in fungi and plants. In this paper, we describe the properties and the analysis of the function of the first PH domain ever reported to occur in a cell wall carbohydrate synthase.

First, we performed a bioinformatic analysis of the PH domain of cellulose synthase 2 from S. monoica (SmCesA2). A Blast search against the proteins in the PDB database (http://www.ncbi.nlm.nih.gov/blast/Blast.cgi) identified the C-terminal PH domain of the TAPP1 human protein (“tandem PH-domain-containing protein-1”; PDB ID: 1EAZ_A) as the closest homolog of the PH domain of SmCesA2 (E-value=2e-05).

The TAPP1 PH domain binds phosphatidylinositol-3,4-biphosphate (PtdIns-(3,4)-P2) with high affinity and specificity (Dowler et al., 2000). All PH domains with similar binding activity are gathered in group 1 (see section 1.11.5). The TAPP1 PH domain is targeted to the plasma membrane and co-localizes with F-actin in B-lymphoma cells (Marshall et al., 2002). This domain and other PH domains from group 1 contain so-called PtdIns-(3,4)-P2 and PtdIns-(3,4,5)-P3 binding motifs (PPBM) characterized by the consensus sequence of Lys-X-Sma-X6-11-Arg/Lys-X-Arg-Hyd-Hyd where X represents any amino acid, Sma corresponds to a small amino acid and Hyd to a hydrophobic amino acid (Isakoff et al., 1998). The PH domain of SmCesA2

(SmCesA2PH) may interact with PtdIns-(3,4)-P2 and/or PtdIns-(3,4,5)-P3 since its sequence contains a PPBM motif. The full length of SmCesA2PH was fused to an N- terminal hexa-histidine tag, expressed in E. coli BL21(DE3) and purified by affinity chromatography to obtain recombinant SmCesA2PH protein that was used to determine its binding specificity in vitro. The identity of the purified recombinant protein was confirmed by mass spectrophotometry.

A protein-lipid dot-blot assay was performed using a so-called “PIP strip” membrane carrying different phosphoinositides and other types of phospholipids, and all samples were blotted at the same concentration. The results showed that SmCesA2PH is promiscuous in contrast to its TAPP1 counterpart. It binds PtdIns-

(3,4)-P2 and PtdIns-(3,4,5)-P3, similar to the PH domain of TAPP1, but it also binds monophosphatidylinositols (PtdIns-(3)-P, PtdIns-(4)-P, PtdIns-(5)-P) and PtdIns-(3,5)-

P2, albeit with a lower affinity. In addition, a weak binding to phosphatidic acid and

60 phosphatidylserine was detected (Figure 26). To complement the first screening, a second type of membrane called “PIP array” was used to obtain more quantitative data with respect to the relative affinity of SmcesA2PH for the different lipids analysed. The results showed that SmCesA2PH binds strongly to almost all phosphorylated lipids tested (Figure 2B, paper II).

Sphingosine-1- LPA

LP PtdIns(3,4)P2

PtdIns(3,5)P2 PtdIns

PtdIns(4,5)P2 PtdIns(3)P

PtdIns(4)P PtdIns(3,4,5)P3

Phosphatidic PtdIns(5)P

PE Phosphatidylserin

PC Blan

Figure 26. Lipid binding activity of SmCesA2PH. LPA, lysophosphatidic acid; LPC, lysophosphocholine; PE, phosphatidylethanolamine; PC, phosphatidylcholine

Some PH domains from mammalian proteins have also been found to interact with other proteins, more specifically cytoskeleton-related proteins. In vitro pull-down assays using F-actin and microtubules were therefore performed to determine if SmCesA2PH is able to bind these cytoskeleton-related proteins (Figure 3A and 3B, paper II). The data showed that after a pre-incubation at room temperature, SmCesA2PH formed sediment together with F-actin and microtubules, thereby confirming its ability to interact with F-actin and polymerized tubulin. These in vitro approaches were complemented by an in vivo localization assay. The assay was performed using human U2OS osteosarcoma cells and SmCesA2PH fused to GFP (C-terminal). SmCesA2PH_GFP was detected in the nucleoplasm, with a higher intensity in the nucleoli (Figure 4I, paper II) and a higher exposure revealed that SmCesA2PH_GFP was also localized to the cytoplasm (Figure 4J and M, paper II).

61 In contrast, the control for cytosolic localization (chloramphenicol acetyltransferase- GFP, (CAT-GFP)) was distributed evenly throughout the cytosol, as expected. Another control was also included in this experiment where GFP was expressed alone (data not shown). Although GFP was found in the nucleus, the CAT-GFP signal was not higher in the nucleolus as compared to the cytoplasm (Figure 4B and E, paper II). Another interesting finding is that F-actin was shown to co-localize with SmCesA2PH_GFP at the distal borders of the cells, when stained with phalloidin, (Figure 4L and O, paper II) but, such localisation was not found for the control (CAT- GFP), (Figure 4D and G, paper II). Thereby this provides further evidence of the specificity of the interaction between SmCesA2PH_GFP and F-actin. A similar approach was used for co-localization studies with microtubules but the data were not conclusive. Further experiments are needed to be performed to confirm the in vivo interactions between SmCesA2PH and microtubules.

3.3 Biochemical and structural characterization of the SmChs1 MIT domain (Paper III)

Both Chs1 and Chs2 from S. monoica contain N-terminal microtubule interacting and trafficking domains (MIT) (Guerriero et al. 2010). MIT domains are known to occur in some mammalian enzymes, but prior to the work of Guerriero et al. (2010) they had never been reported in any enzyme that acts on carbohydrates. Moreover, the presence of MIT domains in carbohydrate synthases seems to be specific to the oomycete chitin synthases.

The full length of Sm_MIT1 was fused in frame with an N-terminal hexahistidine tag in the pET-28a(+) vector as a first step toward the isolation, characterization and structural analysis of the protein by NMR. The protein was expressed in E. coli BL21(DE3) and purified using cobalt affinity chromatography (Figure 1, paper III). Its identity was confirmed by mass spectrometry (MS/MS) analysis after partial trypsin hydrolysis. Sm_MIT2 was cloned and expressed under the same conditions as Sm_MIT1, but the recombinant protein (Sm_MIT2) was systematically precipitating at the concentration required for NMR analysis. Therefore, the structure of Sm_MIT2 could not be determined by NMR. Instead, the structure of the Sm_MIT2 was first

62 modelled using in silico approaches which revealed three alpha helices, as found for Sm_MIT1. The purified recombinant Sm_MIT1 and Sm_MIT2 domains were analysed by CD spectroscopy and both recombinant proteins were properly folded, (Figure 2A, 2B, paper III) and showed similarity to the MIT domain from mammalian proteins that reveal the presence of α-helices in the far UV spectrum (Scott et al., 2005).

Furthermore, CD spectroscopy was also used to study the thermostability of both MIT domains. MIT1 is characterized by a melting temperature (TM) of approximately 56ºC whereas the TM for Sm_MIT2 is of 42 ºC (Figure 2C, paper III). Although both MIT domains are stable at temperatures as high as 40 ºC the absence of a well-defined plateau at low temperatures indicates some possible changes in the helicoidal structure.

Previous research has shown that MIT domain can interact with cytoskeleton proteins in mammalian cells (Berruti, 2010). Therefore, since the Sm_MIT1 and MIT2 recombinant proteins were properly folded and thus expected to be functional in vitro, we investigated whether they were able to interact with polymerized tubulin and F- actin using in vitro assays. Of the two cytoskeletal proteins tested neither had affinity for the MIT1 and MIT2 domains of S. monoica. The absence of interaction of the recombinant Sm_MIT1 and Sm_MIT2 with microtubules and F-actin may be due to the absence of an MTBD domain in the oomycete proteins. Enzymes with MIT domains, like spastin, in fact interact and severe microtubules via a so-called microtubule-binding (MTBD) domain, which is distinct from the MIT domain (White et al., 2007). The function of the latter is poorly understood apart from the fact that it is important for the activity of the enzyme (Yang et al., 2008).

Until recently, no report showing the capacity of MIT domains to bind phosphoinositides was available (Iwaya et al., 2013). We performed in vitro phosphoinositide binding assays and demonstrated that the S. monoica MIT domains bind strongly to phosphatidic acid (PA) and, to a lesser extent, to phosphoinositides (Figure 6A-E, paper III). The high affinity of Sm_MIT1 and Sm_MIT2 for PA is interesting as PA is involved in membrane fusion (Figure 27). Thus, assisted by its strong interaction with PA during membrane fusion, the MIT domain of the oomycete

63 chitin synthase may likely be involved in the transfer of the enzyme from vesicles to the plasma membrane.

Figure 27. Densitometric analysis of Sm_MIT1 and Sm_MIT2 binding on a Membrane ArrayTM, with standard deviations from three independent experiments. AU: arbitrary units.

Applying a different approach, we used the yeast two-hybrid system to study the interaction of the Sm_MIT1 and Sm_MIT2 domains with other proteins. The MIT2 domain was found to bind to the delta (δ) subunit of the adaptor protein AP3 (Table S2, paper III), and other approaches should be used to validate this findings. In yeast and mammals, AP3 is known to bind a dileucine signal in the [D/E]XXXL[L/I] motif of interacting cargo proteins (Darsow et al., 1998; Vowel and Payne, 1998). More precisely, this motif is known to interact with the δ/σ hemicomplex or the σ subunit itself (Janvier et al., 2003). Interestingly, our analyses revealed the occurrence of the dileucine motif in the MIT domain of SmChs2, suggesting its likely involvement in the interaction between the δ large subunit of AP3 and Sm_MIT2. It has been suggested that AP3 is involved in the intracellular targeting of transmembrane glycoproteins to endosomes/lysosomes (Le Borgne et al., 1998). Therefore a possible implication of the MIT domain in the degradation/recycling pathway of chitin synthase might be possible. For Sm_MIT1 the yeast two-hybrid system did not reveal any interacting partners and interestingly there are no dileucine motifs in Sm_MIT 1. Therefore, the oomycete MIT domains may require a dileucine motif to bind to AP proteins similar to the proteins in mammalian and yeast counterpart.

The three-dimensional structure of the recombinant Sm_MIT1 domain was solved by NMR (Figure 28). The structure consists of a bundle of three antiparallel helices,

64 similar to that observed previously for the VPS4A MIT domain (Scott et al. 2005). Helix 1 comprises residues Ile2-Ala15, helix 2 residues Arg20-Lys41 and helix 3 residues Gln45-His71. The third helix is the longest with 26 residues. It shows a tight interaction with the end of the second helix. The Sm_MIT1 structure is similar to other well characterized MIT domains such as for instance those from the human Vps4a and Vps4b proteins. In addition, the NMR structure is consistent with the in silico model of MIT 2 and showed a similar structural and dynamic features.

Figure 28. NMR structure of Sm_MIT1. Yellow spheres show residues with normalized ASA scores <0,10 (the least solvent exposed). Orange shows residues with ASA scores <0,25 (more solvent exposed).

3.4 Modelling of the interactions between the MIT domain of SmChs1 (Sm_MIT1) and lipids (Paper IV)

As described in paper III, the MIT domain of SmChs1 interacts strongly with phosphatidic acid (PA). In this paper, we investigated in detail the types of interactions involved and identified amino acids important for binding with the POPA (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidic acid) bilayer.

The MIT domain can be approximated as cuboidal with 6 faces (P1-P6) (Table III). To determine which orientation allows for the strongest absorption, we performed molecular dynamics (MD) simulation (see Materials & Methods in paper IV). We included two membranes in our study, a lipid bilayer made of POPA and a control composed of POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphatidylcholine).

65 Our simulations indicated orientations P3 and P5 provide the highest binding affinities (Table III). Interestingly, P3 and P5 share the same binding residues (Arg7, Arg11, Arg20 and Arg22) (Table III). These basic residues are capable of both electrostatic and hydrogen bond interactions and consequently have a higher affinity to the POPA membrane. The combination of these two non-covalent interactions is termed “salt-bridge interaction” and the salt-bridge values for P3 and P5 are the highest of all orientations examined. As shown in Figure 7 from paper IV, the binding free energies correlate with the total salt-bridge values for each orientation. This highlights the importance of the arginine residues as arginine has a higher salt-bridge value than lysine. The arginine residues are mostly located in helix 1 and the N- terminal of helix 2 (Figure 29), and consequently this region was described as a “binding hotspot”. Upon closer examination, we find that the binding hotspot is in full contact with the membrane in orientations P3 and P5. In contrast, the binding hotspot is not in full contact with the membrane in orientations P1 and P2, corresponding to lower binding free energy values relative to P3 and P5. Finally, in orientations P4 and P6 the binding hotspot is not in contact with the membrane and as a result, the binding free energy values are very low.

Identical simulation conditions were used to examine the interactions between the MIT domain and the POPC membrane. The highest binding free energy value for MD simulations with the POPC membrane, -12.33 kcal/mol, was far lower than that detected for the POPA membrane (Table 1, Table 2, paper IV). The interactions with the POPC membrane are predominantly composed of the charged MIT residues binding to the phosphate and quaternary ammonium groups by direct charge-charge attractions. The weak binding of MIT residues to the POPC membrane can be

66 Table 3. Study of the different orientations (P1-P6) taken by Sm_MIT1 domain on the POPA membrane before and after adsorption. Number of Binding Free a Binding Salt-bridge Initial State Final State Contact Energy Residues c Value Residues b (kcal/mol)

Arg20, Arg22, P1 15 2.86 -25.59 Lys26, Lys41

Arg7, Arg11, Lys46, P2 20 4.28 -27.89 Arg48, Lys54, Lys56 Arg7, Arg11, P3 14 Arg20, 4.43 -28.63 Arg22, Lys26

Lys41, Lys46, P4 14 2.29 -20.50 Arg48, Lys56

Arg7, Arg11, P5 10 4.00 -28.82 Arg20, Arg22

Lys41, P6 9 Lys46, 1.86 -18.00 Arg48

a In the pictures of intial and final states, MIT is shown in cartoon mode and lipid molecuels in stick mode. Water and ions are not shown for clarity. b Contact residues are those within 3 Å of POPA molecules c Binding residues are those having electrostatic and hydrogen bond interactions with POPA molecules.

Figure 29. Structure and sequence of Sm_MIT1. Basic residues are in blue, acidic in red, polar in green and nonpoplar in white. attributed to the self-saturated state of the membrane surface. The negative phosphate groups and the positive choline groups that constitute the POPC membrane allow for inter-molecule electrostatic interactions, which the MIT domain must compete with in order to bind. Finally, the size of the lipid head group can also affect binding. More specifically, the phosphatidylcholine head group of the POPC membrane is large relative to the phosphate group of the POPA membrane, further contributing to the difference in absorption rate between the two membranes. In short, our analysis has led us to conclude that the electrostatic and spatial differences of POPA and POPC molecules are the fundamental basis for the varied binding affinity of the MIT to these two membranes.

3.5 Characterization of two putative chitin synthases (AeChs1 and AeChs2) from microsomal fractions of Aphanomyces euteiches (Paper V)

The pathogen Aphanomyces euteiches is known to infect the roots of legumes and to contain two putative chitin synthases (Badreddine et al., 2008). Paper V shows for the first time the formation of a soluble and insoluble product from in vitro activity assays using microsomal fractions of A. euteiches. A previous study reported only

68 the presence of GlcNAc-based carbohydrates (Badreddine et al., 2008; Mélida et al., 2012) in the cell wall of the microorganism but no Chs activity was detected in vitro (Badreddine et al., 2008).

Microsomal fractions of A. euteiches were prepared and used for enzymatic assays as described in the Materials and Methods section of Paper V. Interestingly, two types of products were formed and the amount of each product synthesized varied depending on the occurrence of effectors in the assay mixture. For instance, the presence of trypsin in the assay stimulated the synthesis of the water-insoluble product than the soluble product (Figure 1, paper V). Mn2+ increased the synthesis of soluble products whereas insoluble product formation was unaffected by the presence of divalent cations (Figure 3, paper V). Strong inihibition of the formation of insoluble products was observed in the presence of Ni2+ (Figure 3, paper V). Interestingly, the presence of N-acetylated chitooligomers moderately stimulated the synthesis of the insoluble product whereas the amount of soluble product formed in vitro was lower (Figure 4, paper V). The amount of product synthesized is influenced by the degree of polymerization (DP) of the added chitooligomer. As a general observation, a higher DP favoured the synthesis of insoluble products whereas the opposite trend was observed for the soluble product. These differences show that the type of product synthesized by the A. euteiches Chs may be regulated by various effectors.

The hydrolysis of the insoluble product by chitinase confirmed the formation of chitin- like products in the in vitro assay mixtures (Figure 1, paper V). Analysis of the soluble products by thin-layer chromatography (TLC) revealed the formation of mostly GlcNAc dimers (Figure 2, paper V). Unfortunately, the type of linkages could not be determined due to the low amount of disacharide formed. Chitin synthase activity was also strongly inhibited by nikkomycin Z (Figure 1, paper V). In order to further characterize each Chs gene product from A. euteiches, heterologous expressions of AeChs1 and AeChs2 were attempted in different strains of E. coli but no activity was detected in the recombinant proteins preparations.

69 4 Conclusions and perspectives ______

The general objective of this thesis was to better understand oomycete cell wall biosynthesis in order to develop new approaches to tackle infections related to these organisms. A more specific aim was to characterize a pleckstrin homology domain in cellulose synthases and a microtubule interacting and trafficking domain in chitin synthases from Saprolegnia monoica.

In paper I we reported on two assays to determine the activities of glycosyltransferases (GTs). The radiometric assay allows for a fast and precise method to biochemically characterize GTs by using membrane fractions and avoiding long purification steps that may increase the risk of losing enzyme activity. The method was used in paper V to investigate Chs activity from microsomal fractions of Aphanomyces euteiches. Insoluble chitin-like products and soluble chitobiose were detected in the in vitro reaction mixtures. The activity assays suggest that Aphanomyces euteiches Chs1 and Chs2 are active and that the presence of chitooligosaccharides stimulated the formation of insoluble products but inhibited the accumulation of soluble products.

Paper II showed the unique presence of PH domains in oomycetes cellulose synthases and provided information about their binding activity. Like other characterized PH domains, the SmCesA2PH domain showed promiscuous binding to phosphatidylinositol. To better understand these interactions the structure of SmCesA2 in the presence of PtdIns ligand is needed. The C-terminal PH domain of the human protein TAPP1 was identified to be the most similar to the SmCesA2PH domain. The SmCesA2PH domain did not bind to PtdIns(3,4)P2 with the same affinity and specificity as does the TAPP1 PH domain, but it interacted with F-actin under in vitro conditions and co-localized in vivo with F-actin. Interestingly, SmCesA2PH did bind in vitro to microtubules, which is not the case for TAPP1 and other PH domains. The binding to cytoskeleton proteins suggested that the PH domain could be involved in the targeting of cellulose synthases to the plasma membrane.

70 Papers III and IV described the properties of the MIT domains of chitin synthases from S. monoica. The structure of Sm_MIT1 was modelled and experimentally solved by NMR. The simulation study predicted a higher affinity for phosphatidic acid compared to phosphatidylcholine due to electrostatic interactions and hydrogen bond. Arginine residues located in helix 1 and the N-terminal of helix 2 were found in silico to be key to Sm_MIT1 interaction with POPA. The interactions between the MIT domain and phosphatidic acid might be essential to facilitate the fusion of vesicles containing Chs at the plasma membrane. In addition, the interactions of the MIT domains with PtdIns suggest involvement in cellular targeting processes. In our studies on the S. monoica MIT domains, we did not find any in vitro interaction with microtubules or F-actin. However, similar to the Ustilago maydis myosin chitin synthase 1 (UmMcs1), motor proteins like kinesin and myosin could be involved in the transport of vesicles containing the oomycete Chs. These interactions could stabilize and tether the vesicles, and thereby increase the residence time at the plasma membrane, to promote fusion. Another interesting discovery is the interaction between Sm_MIT2 and the AP3 δ subunit of S. monoica. To better visualize the different types of interactions that Sm_MIT2 could be involved in, I propose a hypothetical model that suggests different pathways that could be used by Sm_Chs2 to reach the PM. The subsequent recycling and degradation of the enzyme are also represented in the model (Figure 30).

Figure 30. Hypothetical model showing intracellular trafficking of chs in hyphae and possible targeting pathways of Saprolegnia monoica chitin synthases to the plasma membrane (PM) are represented. One of the trafficking pathways to the PM involves the participation of the cytoskeleton (black arrows) whereas the other proposes the migration of vesicles containing Chs from the trans-Golgi to the PM in an, independent manner (brown arrow). The AP-3 putative functional pathway as described in yeast and mammalian cells is represented in green arrows. The pathway from trans-Golgi to endosomes

72 could allow the formation of a storage compartment of SmChs2 at the endosome that supplies the plasma membrane, as proposed by Starr (2012) for ScChs3. The blue frame shows possible pathways for the retargeting/recycling of Chs by the AP-2, a protein involved in endocytosis. One arrow (blue) shows retargeting/recycling of Chs from endosome to PM, whereas the other blue arrow from endosome to TGN showing Chs can be recycled for re-entry back into the secretory pathways (Feyder et al., 2015).

The Golgi membrane is enriched in PtdIns(4)P. The interaction between PtdIns(4)P and the MIT domain might stabilize the soluble part of the Chs2 during the targeting process. Moreover, experiments in yeast showed that PtdIns(4)P can act as a signal to trigger the transport of ScChs3 from the Golgi apparatus to the PM (Schorr et al., 2001). The integration at the PM is believed to be facilitated by the interaction of the MIT domain with PA.

Although we cannot yet confirm the function of the PH and MIT domains in GTs, our work has allowed for a better understanding of their possible functions.

Efforts were made to get the structure of SmCesA2PH by expressing different truncated sequences of the domain, but no protein crystals have been obtained so far. The work on solving the structure should continue in order to better understand the interactions between cytoskeleton proteins and PtdIns. Further work to develop an in vivo binding assay with microtubules can help to confirm whether microtubules have a role in the delivery of SmCesA2 to the membrane.

The research on MIT domains will be completed by NMR to investigate whether any changes occur in the MIT domain structure in the presence of phospholipids. This would confirm which PtdIns are interacting with the MIT domain and allow for identification of amino acid residues that are involved in the binding. The interaction between Sm_MIT2 and the AP3 δ subunit will be further investigated. An in vitro binding assay with the AP3 δ subunit and Sm_MIT2 could confirm the interaction that was discovered in the yeast two-hybrid assay. If these interactions are verified, point mutations in the dileucine-based motif of MIT2 could be performed to identify the key amino acids involved. If possible, NMR could also be used to better characterize these interactions. Finally, it would also be interesting to verify if AP2 is interacting with the MIT domain because it is known to bind the dileucine-based motif. This

73 would demonstrate the participation of the MIT domain in the endocytosis process of Chs.

Attempt was made to determine the in vivo localization of Chs1 and Chs2 from S. monoica via transient expression of full-length and truncated gene constructs in Nicotiana benthamiana using the agroinfiltration technique. Results were inconclusive and similar experiment could be repeated in future in both N. benthamiana and ideally in S. monoica if an efficient transformation protocol for oomycete can be developed. As S. cerevisiae chitin synthases do not contain MIT domains, the yeast system could therefore be a useful heterologous expression host to study the trafficking of oomycete Chs. A better understanding of cell wall formation in oomycete could lead to new ways of tackling the growth of these pathogens for disease control.

74 5 Acknowledgements ______

The opportunity to pursue my PhD came while I was doing my diploma work supervised by Prof. Tuula Teeri and Assoc. Prof. Emma Master. Both have been a great inspiration for continuing my research career.

It has been a challenging stage to get my PhD and many people have been involved throughout these years. I am grateful to all of you for your help and advice. Here, I wish to express my special gratitude to all of you and, I hope, I am not forgetting anyone.

I would like to express my deepest appreciation to both Prof. Tuula Teeri and Prof. Vincent Bulone for their time, patience and excellent guidance throughout my PhD. Special thanks to Prof. Vincent Bulone, I have enjoyed working with you and without you, my thesis would not have been possible.

I would like to thank all my co-authors: Dr. Lars Arvestad, Dr. Henrik Aspeborg, Prof. Lars Berglund, Dr. Kristina Blomqvist, Dr. Arnaud Bottin, Prof. Vincent Bulone, Dr. Nuria Butchosa, Assoc. Prof. Christina Divne, Assoc. Prof. Ines Ezcurra, Dr Johanna Fugelstad, Dr. Gea Guerriero, Prof. Sophia Hober, Elvira Hukasova, Guanglin Kuang, Dr. Manoj Kumar, Felicia Leijon, Dr. Thomas Larsson, Dr. Lijun Liang, Dr. Arne Lindqvist, Prof. Lena Mäler, Dr. Lauren S. McKee, Assoc. Prof. Ewa Mellerowicz, Dr. Hugo Melida, Dr. Podjamas Pansri, Dr. Alex Rajangam, Dr. Vaibhav Srivastava, Prof. Bjorn Sundberg, Dr. Gustav Sundqvist, Scarlett Szpryngiel, Prof. Tuula Teeri, Dr. Yaoquan Tu, Weihua Ye and Assoc. Prof. Qi Zhou for your great work to achieve our common goals.

I would like to acknowledge Annie Inman, for helping with additional correction of my thesis, articles and Assoc. Prof. Qi Zhou for critical reading of my thesis.

I would also like to thank our present and previous administrative staff, Nina Bauer, Marlene Johnsson and Lotta Rosenfeldt for your great help throughout these years. Thanks also to Ela Nilsson, for taking care of the lab and Kaj Kauko for IT support and update on the latest news about cell phone and computer technology :)

75 Many people have been working with me on floor 2 during my PhD and I wish to say thank you to everybody, especially the people in Bulone’s group for the great atmosphere and discussions. The list of people would be never-ending  but I would like to mention Dr. Jens Eklof, Dr. Farid Ibatullin and Dr. Per-Olof Syrén for sharing long evenings and weekends working in the lab.

Special thanks to my office mates Dr. Núria Butchosa (and her lovely orchids:), Paul Dahlin, Stefan Klinter, Felicia Leijon Dr. Erik Malm, Zhili Pang, Ela Rzeszutek (for eating my cookies and chocolate with happiness), and Dr. Hwei-Ting Tan for the nice chats and making the work even more enjoyable. Thanks Ela R. for your interesting questions and scientific discussions and Peter Hendil-Forssell, Dr. Cortwa Hooijmaijers, Dr. Ibatullin, Dr Svetlana Rezinciuc and Dr. Hwei-Ting Tan for technical help. I also wish to say thanks to the people from Industrial Biotechnology, especially Dr. Johan Jarmander for your tips and tricks in finalising my thesis. To my former officemates from the wood biotechnology Dr. Alex Rajangam and Dr. Anders Winzél, thank you for the great time, help and friendships.

I would also like to thank Doc. Henrik Aspeborg, Doc. Torkel Berglund, Prof. Kalle Hult, and Doc. Anna Ohlsson for sharing their passion for nature with me.

Last but not least, my special thanks go to my family and my girlfriend, Judith for their support and encouragements throughout all these years away from home. Despite the distance from home, my family members are always close to my heart.

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