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
© Christian Brown, 2015
School of Biotechnology
Royal Institute of Technology
AlbaNova University Center
106 91 Stockholm
Sweden
Printed by Universitetsservice US-AB
Drottning Kristinas väg 53B
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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
© Christian Brown, 2015
i
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 (PHdomänen) 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 CHSproteiner. 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)
ii
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 microtubuleassociated 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
iv
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
vi
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).
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 (NovaesLedieu, 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).
A
B
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 Nacetylglucosaminyl (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-GlcNAcbased 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 βglucan 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-1,6glucans 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).
12
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 carbohydrateactive 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 GTA 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).
14
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).
15
Figure 9. Proposed mechanism for retaining glycosyltransferases (adapted from Yip and Withers,
2+
2004). Divalent cations (M ) 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 αchitin 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).
19
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 todate, 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 GPIanchored 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).
23
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; RuizHerrera and Bartnicki-Garcia 1976), Aspergillus flavus, Aspergillus nidulans (LopezRomero 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).
32
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 UDPGlc 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 porinlike 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.
37
B
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 (DíazMoreno 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 “tyrosinebased” 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 YXXΦ 2 and the NPXY 3 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).
40
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 (NewellLitwa 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 MITinteracting 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 ESCRTIII 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 (RollMecak 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 deubiquitinating 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 phosphoinositidebinding 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.
50
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).
51
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 proteinprotein 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 domaincontaining 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 (MartínezRucobo 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
14
C or
3
H) 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,
13
C-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 Nterminal 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
PtdIns(3,4)P2
LP
PtdIns(3,5)P2
PtdIns
PtdIns(4,5)P2
PtdIns(3)P
Figure
26.
Lipid
PtdIns(4)P
PtdIns(3,4,5)P3
PtdIns(5)P
Phosphatidic
binding
PE
Phosphatidylserin
PC
Blan
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 acetyltransferaseGFP, (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 (CATGFP), (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 Factin 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.
TM
Figure 27. Densitometric analysis of Sm_MIT1 and Sm_MIT2 binding on a Membrane Array , 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 Nterminal 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
Contact
Residues b
Binding
Residues c
Salt-bridge
Value
Binding Free
Energy
(kcal/mol)
15
Arg20,
Arg22,
Lys26,
Lys41
2.86
-25.59
20
Arg7,
Arg11,
Lys46,
Arg48,
Lys54,
Lys56
4.28
-27.89
14
Arg7,
Arg11,
Arg20,
Arg22,
Lys26
4.43
-28.63
14
Lys41,
Lys46,
Arg48,
Lys56
2.29
-20.50
P5
10
Arg7,
Arg11,
Arg20,
Arg22
4.00
-28.82
P6
9
Lys41,
Lys46,
Arg48
1.86
-18.00
Initial State a
Final State
P1
P2
P3
P4
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.
67
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 chitinlike 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).
71
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.
76
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