Identification and Functional Characterization of Core Effectors from Cyst Nematodes Through Comparative Genomics

Institut für Nutzpflanzenwissenschaften und Ressourcenschutz

Molekulare Phytomedizin

Identification and functional characterization of core effectors from cyst nematodes through comparative genomics

Dissertation

zur

Erlangung des Grades

Doktor der Agrarwissenschaften (Dr. agr.)

der

Landwirtschaftlichen Fakultät

der

Rheinischen Friedrich-Wilhelms-Universität Bonn

vorgelegt von

Somnath Suresh Pokhare

aus

Indien

Bonn, 2018

Referent: Prof. Dr. Florian M.W. Grundler

Gutachter: Prof. Dr. John Jones

Tag der mündlichen Prüfung: 03.12.2018

Angefertigt mit Genehmigung der Landwirtschaftlichen Fakultät der Universität Bonn

This work is dedicated to my beloved wife Devata for her patience and unconditional support during all these years and to our princess Mugdha…!!!

Table of contents

Abstract...... i

Zusammenfassung...... iii

List of figures...... v

List of tables………………………...... ………...vii

Chapter 1: Introduction…………………...... 1

1.1 Nematode diversity and distribution...... 1

1.2 General body structure…….…………………...... 2

1.3 Nematode feeding groups...... 3

1.4 Plant Parasitic Nematodes (PPNs)...... 5

1.5 Cyst nematodes...... 7

1.6 General life cycle of cyst nematodes...... 8

1.7 Syncytia morphology and physiology...... 10

1.8 Nematode effectors...... 12

1.9 Identification of nematode effectors…………………………...... …….....…….....14

1.10 Functional validation of effectors...... 15

1.11 Classification of nematode effectors...... 17

1.12 Comparative genomics...... 21

1.13 Aims of the thesis...... 22

1.14 References...... 23

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel…....………….……39

2.1 Summary…………………………………………………………...….…………….…40

2.2 Introduction…………………………….……………………….…………………...…41

2.3 Material and methods....………………………………………………....……….…….43

2.4 Results…………………………….……………………………….…...………………45

2.5 Discussion……………………….………………….……………….…………………46

2.6 Figures………………………….………………………………………...……………48

Table of contents

2.7 References……………………………………………………………………………..50

Chapter 3: Signatures of adaptation to a monocot host in the plant-parasitic nematode Heterodera sacchari…………..…………………………………………………....……53

3.1 Abstract………....…………………………………………………………………...…54

3.2 Introduction…………….……………………………………………………...... ….….55

3.3 Materials and methods.……………………………………….…………..……………57

3.4 Results and discussion.……………………………………………...... …………….…62

3.5 Conclusion……………………………………………………………...……………...66

3.6 Tables...……………………………………………………………..………………….68

3.7 Figures...... 69

3.8 References………………………………………..…………….………………………74

Chapter 4: General conclusion……………………………………..………………………84

4.1 Supplementary figure…..………………………………………………………………94

4.2 References……………………………………………………..…………….…………94

Acknowledgement…………….………………………………………...……………..……102

Abstract

Plant parasitic nematodes (PPNs), are one of the most important constraints in agricultural production causing substantial losses in quantity and quality of food. The economically most important PPNs are the sedentary endoparasitic root-knot and cyst nematodes. Heterodera sacchari, a sedentary endoparasitic cyst nematode, is an important parasite of sugarcane and rice crops and is an emerging threat to intensive upland rice cultivation especially in west Africa. Like other species of cyst nematodes, pre-parasitic infective juveniles (J2s) of H. sacchari penetrate the host roots and induce a metabolically active syncytial feeding cell system in the stele. To gain more detailed insights into the H. sacchari-rice interaction, we established an in vitro culture system for H. sacchari on Nipponbare rice in pluronic gel. We confirmed that unlike other cyst nematodes, H. sacchari reproduces by parthenogenesis and males are rare except under high stress conditions. At 25 days post inoculation (dpi), developing females start to produce eggs inside the body. At around 40 dpi, the cuticle of the females started to tan, and the females became darker as the life cycle progressed. We found that H. sacchari can successfully complete its life cycle in 7-9 weeks at 25 °C on Nipponbare rice in pluronic gel. Results of hatching assays showed that hatching of juveniles from eggs contained in cysts was stimulated by 3mM ZnCl2 but not by rice root exudates. The established in vitro culture system can be efficiently used to collect post-infective stages of nematode and host tissue at different time points for various studies and also to screen different rice cultivars for nematode resistance/susceptibility. Using in vitro material, we generated transcriptome data from two different life stages (pre-parasitic J2s and 15 days post-infection) using Illumina MiSeq next-generation sequencing with a total of 17,086,132 paired-end 2x 250 bp reads.

A multi-gene phylogenetic analysis using Core Eukaryotic Gene Mapping Approach (CEGMA) genes from a range of nematodes showed that H. sacchari and the cereal cyst nematode H. avenae (both monocot parasites) evolved from a common dicot-parasitic ancestor. Comparisons of putative effector and non-effector genes from H. sacchari to their closest homologues in potato cyst nematode G. rostochiensis showed that effector genes are more divergent than non-effector genes.

We found some effectors with conserved expression profile and likely function (e.g. cellulase, chorismate mutase, 19C07). Transcripts encoding genes similar to the SPRYSEC family of effectors were also present in H. sacchari dataset, suggesting conservation of this effector family across cyst nematodes. The transcriptome of H. sacchari contains seven transcripts that encode proteins similar to CLE effectors. The CLE like peptides encoded by

Abstract

H. sacchari are more similar to rice CLEs than any other plant or nematode CLEs. We also demonstrated that exogenous application of H. sacchari CLE peptide induced a short root phenotype in rice while peptides from the dicot parasite H. glycines did not produce any phenotype. These results provide insights in effector evolution that co-occurred with the transition from a dicot-parasitic to a monocot-parasitic lifestyle.

Zusammenfassung

Pflanzenparasitäre Nematoden verursachen substantielle Verluste in der Quantität und der Qualität von Feldfrüchten und stellen damit ein großes Problem in der Landwirtschaft dar. Die ökonomisch wichtigsten PPNs sind sedentäre endoparasitische Wurzelgallen und Zystennematoden. Heterodera sacchari, ein sedentärer endoparasitischer Zystennematode, ist ein wichtiger Schaderreger in Zuckerrohr und Reis. Er gefährdet insbesondere den zunehmend intensiven Reisanbau in West Afrika. Wie bei anderen Arten von Zystennematoden dringen präparasitische Juvenile (J2s) von H. sacchari in die Wurzel der Wirtspflanze ein und induzieren ein metabolisch aktives, synzytiales Nährzellsystem im Leitgewebe. Um detaillierte Einblicke in die Interaktion zwischen H. sacchari und Reispflanzen zu gewinnen, etablierten wir eine in vitro Kultur von H. sacchari an der Reissorte Nipponbare in Pluronic Gel. Wir konnten bestätigen, dass sich H. sacchari, im Gegensatz zu anderen Zystennematodenarten, durch Parthenogenese fortpflanzt und dass Männchen nur unter Stressbedingungen gehäuft vorkommen. Weibchen begannen mit der Eiproduktion 25 Tage nach der Inokulation. Nach 40 Tagen färbte sich die Kutikula der Weibchen dunkler. Wir beobachteten, dass H. sacchari in Nipponbare bei 25°C in sieben bis neun Wochen den Lebenszyklus erfolgreich durchläuft. Die Ergebnisse von Schlupfstudien zeigten, dass der Schlupf von Juvenilen aus Zysten durch 3mM ZnCl2 stimuliert wird, aber nicht durch Wurzelexsudate. Die etablierte in vitro Kultur kann effizient für die Sammlung von post-infektiven Stadien, infiziertem Wurzelgewebe sowie für das Screening von Reiskultivaren verwendet werden. Unter Verwendung von in vitro Proben erhoben wir Transkriptomdaten von zwei Stadien (präparasitische J2 und parasitische Juvenile 15 Tage nach Inokulation) anhand von Illumina MiSeq Next-Generation-Sequenzierung, die sich insgesamt auf 17.086.132 paired-end 2x 250 bp Reads belaufen.

Eine phylogenetische Analyse mit dem Core Eukaryotic Gene Mapping Approach (CEGMA) mit Genen von verschiedenen Nematodenarten ergab, dass H. sacchari und der Getreidezystennematode H. avenae (beide Parasiten von Monokotyledonen) aus einem gemeinsamen Vorfahren, der Dikotyledonen parasitierte, hervorgegangen sind. Der Vergleich von putativen Effektoren- und Nicht-Effektoren-Genen von H. sacchari mit den nächsten Homologen des Kartoffelzystennematoden G. rostochiensis zeigte, dass Effektoren-Gene stärker voneinander abweichen als Nicht-Effektoren-Gene.

Wir untersuchten einige Effektoren mit konservierten Expressionsprofilen und ähnlichen Funktionen (z. B. Zellulasen, Chorismat-Mutase, 19C07). Transkripte mit Ähnlichkeit zu

iii

Zusammenfassung

Genen der SPRYSEC Effektor-Familie konnten auch bei H. sacchari gefunden werden. Diese Ergebnisse deuten auf eine Konservierung dieser Effektoren-Familie in Zystennematoden hin. Das Transkriptom von H. sacchari beinhaltet weiterhin sieben Transkripte, die für Proteine mit Ähnlichkeit zu CLE Effektoren kodieren. Eine genauere Analyse zeigte, dass CLE Peptide von H. sacchari mehr Übereinstimmung mit Reis CLEs als mit CLE Sequenzen von anderen Pflanzen oder Nematoden aufweisen. Wir konnten außerdem zeigen, dass die exogene Anwendung von H. sacchari CLE Peptiden einen Phänotyp mit kurzen Wurzeln hervorrief. Peptide des Sojazystennematoden H. glycines, einem Parasiten von Dikotyledonen verursachten dagegen keinen Phänotyp. Die Ergebnisse der Arbeit bestätigen den Evolutionsverlauf von Effektoren, der gleichzeitig mit dem Übergang von Dikotyledonen zu Monokotyledonen als Wirtspflanzen erfolgte.

List of figures

Figure 1.1: Nematode distribution with habitat……………………………….....…...... 1

Figure 1.2: Nematode anatomy…………………………………………..…...... 2

Figure 1.3: Nematode feeding groups based on the structure of their mouthparts……..……3

Figure 1.4: A drawing of root cross section showing feeding cell formation by sedentary nematode females………………………………………………….……..……...7

Figure1.5: Typical life cycle of an amphimictic cyst nematode H. schachtii……….……...9

Figure 1.6: Syncytium formed by the soybean cyst nematode, H. glycines in soybean root……………………………………………………………………………..11

Figure 1.7: Illustration of anterior part of juvenile and developing female…………...... 13

Figure 2.1: Levels of susceptibility of different rice varieties to H. sacchari under greenhouse conditions...………………....……………………………………..48

Figure 2.2: H. sacchari hatching assay using water, 3mM ZnCl2 and rice root exudates….48

Figure 2.3: In vitro life cycle of H. sacchari on pluronic gel……………….………...... …49

Figure 3.1: Phylogenetic analysis of 101 CEGMA genes conserved in 18 species of nematodes...…………………………………………………………………….69

Figure 3.2: Analysis of expression patterns of cellulase and chorismate mutase identified in the H. sacchari transcriptome……………………………...... …….………….69

Figure 3.3: Histograms showing distributions of percentage identity between H. sacchari effectors and non-effectors with their homologues in G. rostochiensis.……....70

Figure 3.4: Analysis of expression patterns of two putative SPRYSEC-encoding transcripts of H. sacchari…………………………………..………………………………71

Figure 3.5: Hyper-variable extracellular effector gene family (Hyp) identified from H. sacchari………………………………...……………………………………....71

Figure 3.6: Alignment of CLE-effector-like sequences of H. sacchari...…………….……72

Figure 3.7: CLAVATA3/EMBRYO SURROUNDING REGION-like (CLE) effectors identified from H. sacchari…………………………………………………….72

Figure 3.8: Functional adaptation of H. sacchari CLE effectors to monocot hosts………..73

List of figures

Figure 4.S1: Transient expression of H. sacchari 19C07 effector in the leaves of N. benthamiana showed its localization in the cell periphery….………………....94

List of tables

Table 3.1: H. sacchari sequences similar to putative cell wall degrading and modifying proteins………………………………………………………………...... …..68

Table 3.S1: Primers used for In situ hybridisation...... 68 Table 3.S2: Primers used for quantitative PCR (qRT-PCR)…………………………..……68

vii

Chapter 1: Introduction

1.1 Nematode diversity and distribution

Nematodes, which are commonly called threadworms or roundworms, are one of the largest and most diverse phyla of the animal kingdom. The word nematode comes from the Greek words nemat- meaning thread, and -odes meaning like or resembling. Nematodes are ubiquitous and occupy almost all known habitats, ranging from glaciers to deserts and from hill tops to the bottom of the ocean (Figure 1.1) (Hodda, 2011). They are essentially aquatic animals as they require water for their survival and locomotion, whether this is the aqueous environment of the ocean, a river or a plant or animal host or the thin film of water present within soil matrices. Most nematode species are free-living either in soil or in water (fresh or sea water) that feed on bacteria, fungi even on other nematodes. It is thought that the highest nematode biodiversity is to be found in marine ecosystems. In general, nematode distribution is patchy, as active dispersion is very slow and for short distance only. A handful of soil may contain several thousands of nematodes. Long-distance nematode dispersion is mainly passive and takes place with help of environmental factors (wind and water), anthropogenic activities and also, in some cases with the aid of vectors. Of all the nematodes characterised to date, about 50% nematodes have a marine habitat. The majority of terrestrial nematode species (~25%) are free living while a small proportion of nematode species parasitize plants (~10%) or are parasites of animals including humans (~15%) (Ayoub, 1980; Maggenti, 1981).

10% Marine 15% Free living 50% Animal Parasites 25% Plant parasites

Figure 1.1: Nematode distribution with habitat

Approximately 27000 species of nematode have been described to date but it is estimated that there are between one and ten million species on earth (Quist et al., 2015; Blaxter, 2011;

Chapter 1: Introduction

Zhang, 2013). Little is known about the evolution of nematodes; being soft bodied animals a fossil record is lacking, but it is thought that nematodes are of marine origin (Van Megen et al., 2009; Blaxter and Koutsovoulos, 2015).

1.2 General body structure

Nematodes are usually very small multi-cellular, simple, colourless and, in most cases, transparent animals. Their body shape is like a tiny thread, long and narrow with tapering on both sides in most cases. Nematodes are triploblastic animals i.e. the body is derived from three germ layers of cells during embryonic development: the outmost ectoderm and innermost endoderm having a mesoderm in between. The nematode body cavity is called the pseudocoelom (pseudo-false, coelom-body cavity) as they a lack true body cavity, instead the space between mesoderm and endoderm forms their body cavity (Figure 1.2). The outermost part of the body is the epidermis, or hypodermis, which secretes the cuticle. This is a complex, multi-layered structure formed mainly of collagens and cuticulins that protects the nematodes from the external environment and allows high internal body pressure to be maintained. This cuticle is periodically shed as the nematode grows into adulthood; all nematodes develop through four of these moults during development. The anterior end starts with the head region, which consists of the mouthparts and pharynx connected to the intestine. The anus is the posterior opening of the digestive tract in the tail. In spite of their microscopic size, nematodes possess many of the major organs of higher animals including the intestine, reproductive tissues and nervous system. While the majority of nematode species are microscopic, some animal parasitic species are large enough to be seen with the naked eye. However, the small size of most nematodes means that they are largely unknown to the general public.

Figure 1.2: Nematode anatomy (Source: University of Illinois).

Chapter 1: Introduction

1.3 Nematode feeding groups

Nematodes have a wide range of feeding strategies and show adaptations of the mouthparts to reflect this (Figure 1.3). Bacterial feeding nematodes (a) possess a stoma which is a hollow tube-like structure for bacterial ingestion. One bacterial feeding nematode, Caenorhabditis elegans was chosen as a model system for genetic studies in animals and there are now more genetic and genomic resources available for this organism than for any other animal (The C. elegans Sequencing Consortium, 1998; Kumar et al., 2012). In the case of fungivores nematodes (b), the stoma contains a small stylet without knobs; a needle like structure used to puncture fungal hyphae. Both groups of nematodes play a role in decomposition of organic matter. Plant parasitic nematodes (c), also have a hollow protractible stylet which can be used to puncture the plant cell, to deliver nematode secretions into the host and ingest plant cell contents. Predatory nematodes (d), which feed on other nematodes, possess a tooth like structure, which aids in trapping and tearing the body of prey. Omnivorous nematodes (e), can feed on various food sources depending on the food availability and surrounding environment (Ugarte and Zaborski, 2014).

Figure 1.3: Nematode feeding groups based on the structure of their mouthparts. Bacterial feeder (a), fungal feeder (b), plant feeder (c), predator (d) and omnivore (e) (Ugarte and Zaborski, 2014).

Free living nematodes

Free living nematodes play important role in decomposition and nutrient recycling. In addition, they can be used as biological indicators to analyse soil ecosystem health (Ritz and Trudgill, 1999). The first free-living nematode Turbatrix aceti (vinegar eelworm) was discovered by Borellus in 1656.

Chapter 1: Introduction

Parasitic nematodes

Many described nematodes are parasites of humans, crops and livestock. These nematodes can be divided into animal parasitic nematodes (APN) and plant parasitic nematodes (PPN).

Animal Parasitic Nematodes (APNs)

Nematodes parasitize almost all animals on earth. Some are larger in size than free-living or plant-parasitic species allowing them to be seen easily with the naked eye. The largest APN reported is Placentonema gigantissima, which can reach up to 30 feet long and is a parasite of the sperm whale placenta (Gubanov, 1951). Common animal parasites include intestinal worms (Ascaris and Oxyuris spp.); hookworms (Ancylostoma spp.) and lungworms (Metastrongylus spp.). One gastrointestinal nematode Haemonchus contortus is the most abundant infectious agents in small ruminants (sheep and goat) (Roeber et al., 2013).

Human beings are also widely attacked by nematodes. Rural communities in tropical equatorial regions with insufficient sanitation are more prone to nematode infection. Although 342 species of nematodes found to be associated with humans relatively few are of significant economic importance (Crompton, 1999). The filarial nematodes, Wuchereria bancrofti and Brugia malayi are spread through mosquito bites and cause elephantiasis disease. Currently 120 million peoples from 81 countries carry this infection of which 40 million have disfigurement due to elephantiasis. Another filarial worm Onchocerca volvulus causes onchocerciasis or river blindness, a widespread disease endemic to 27 countries in tropical Africa. About 270,000 individuals are reported to be blind and another 500,000 are thought to have developed a severe visual disability as a result of onchocerciasis (Guerrant et al., 2011). Two hookworm species, Ancylostoma duodenale and Necator americanus and the intestinal roundworm Ascaris lumbricoides are responsible for 125,000 deaths per year (Stepek et al., 2006). It has been calculated that up to 3.5 billion humans carry gastrointestinal nematode infection (Kiontke and Fitch, 2013). Some animal parasitic nematodes can be beneficial. Nematodes of the genera Steinernema and Heterorhabditis are known as entomopathogenic nematodes (EPNs) and parasitize a wide range of insects. These nematodes are widely used as biological control agents for agricultural insect pests (Lacey and Georgis, 2012).

Chapter 1: Introduction

1.4 Plant Parasitic Nematodes

Plant parasitic nematodes (PPNs) are one of the most important constraints in agricultural production and cause substantial losses in quantity and quality of food. Almost every crop is infested by one or more species of PPNs.

The first recording of a PPN was made by Needham, who reported Anguina tritici in the cockles of wheat seeds in 1743. 112 years later, Miles Berkeley discovered root knot nematode (RKN) in 1855 causing galls on cucumber roots grown in a greenhouse in England. In 1859, H. Schacht, a botanist by profession from Bonn, Germany discovered the cyst nematode responsible for beet sickness disease. Later Schmidt (1871), named this nematode Heterodera schachtii. These important discoveries were followed by extensive work in the early 1900s, by Nathan A. Cobb, who is considered as the father of American nematology. Cobb identified around 1,000 nematode species including plant, animal parasites and free living forms (Huettel and Golden, 1991). He was very keen to develop various tools and apparatus for nematological research that led to advancement in the science of nematology. Tools and techniques like fixation and preservation methods, metal mounting slide, flotation and decanting method for nematode extraction from soil developed as a result of his work are still commonly used today (Huettel and Golden, 1991).

The ability to parasitise plants has evolved independently on at least four occasions in the nematodes; the orders Tylenchida, Aphelenchida, Dorylaimida and Triplonchida contain nematodes that can parasitize plants. Around 4100 species of PPNs have been described so far (Decraemer and Hunt, 2006).

Jones et al. (2013), described the most important (top 10) plant parasitic nematode genera. These are RKN (Meloidogyne spp.); cyst nematodes (Heterodera and Globodera spp.); root lesion nematode (Pratylenchus spp.); the burrowing nematode Radopholus similis; the stem and bulb nematode Ditylenchus dipsaci; the pine wilt nematode Bursaphelenchus xylophilus; the reniform nematode Rotylenchulus reniformis; the dagger nematode Xiphinema index; the false root-knot nematode Nacobbus aberrans and the foliar nematode Aphelenchoides besseyi (Jones et al., 2013). General symptoms caused due to nematode attack are not specific and are frequently attributed by growers to nutrient deficiency. Together with other pathogens, nematodes represent a serious threat to global food security. On an average, annual losses caused by PPNs in life sustaining crops are about 10%; while in economically important crops it is 14% (Sasser and Freckman, 1987). It is difficult to accurately calculate losses

Chapter 1: Introduction

caused by PPNs in monetary terms but estimates vary between US$80 billion and US$ 173 billion per year (Nicol et al., 2011; Elling, 2013).

Each plant/crop species is infected by one or more species of PPNs. Plant parasitic nematodes display a range of different host-parasite interactions and attack almost every part (below and above ground) of the plant. The underground parasites can be classified based on their feeding habits. The ectoparasitic nematodes usually have a long stylet and feed on epidermal, outer cuticular cells of the root or root hairs and remain outside the root in the soil (Rehman et al., 2016). These ectoparasites are of minor importance in economic terms, except for those that transmit viruses (e.g. Xiphinema, Longidorus and Trichodorus spp.). The semi endoparasites penetrate several layers of root and feed on cell cytoplasm by inserting the anterior portion of their body. The citrus nematode Tylenchulus semipenetrans and the reniform nematode Rotylenchulus reniformis fit into this class. The endoparasites, which feed on their host while being completely located inside the root, can be subdivided into migratory endoparasites, which move inside the roots while feeding and sedentary endoparasites, which are stationary while feeding. The borrowing nematode Radopholus similis; lesion nematode Pratylenchus spp. and rice root nematode Hirschmaniella spp. are migratory endoparasites (Williamson and Hussey, 1996; Jones et al., 2013). The sedentary nematodes can establish the feeding structure inside host roots (Figure 1.4).

Among all the PPNs, the largest economic losses are caused by sedentary endoparasites i.e. root-knot nematodes (RKN) (Meloidogyne spp.) and cyst nematodes (Globodera and Heterodera spp.) and both these genera fall within the order Tylenchida (Williamson and Gleason, 2003). Both RKN and cyst nematodes are highly evolved obligate biotrophic plant parasites that feed only on cytoplasm of living host cells. These nematodes have evolved the ability to manipulate plant cell fate to form very sophisticated and complex feeding structures inside host roots. Although feeding structures of root-knot and cyst nematodes serve the common purpose of providing food for the completion of nematode life cycle, the process of formation of this novel structure is entirely different and has evolved independently in these two groups (de Almeida-Engler et al., 2011; Grundler and Hofmann, 2011; Bartlem et al., 2014). The feeding structure of RKNs is called the ‘giant cell’ while that of cyst nematodes is called the ‘syncytium’ (Figure 1.4).

The RKN genus constitutes around 98 species (Jones et al., 2013). Many RKN have a very wide host range and are particularly important in hot climates (tropical, subtropical and

Chapter 1: Introduction

Mediterranean region (Saucet et al., 2016). In contrast, cyst nematodes tend to be more host specific and the most damaging species to agriculture are found in temperate or cooler climates (Cotton et al., 2014).

Figure 1.4: A drawing of root cross section showing feeding cell formation by sedentary nematode females (Giant cell of Meloidogyne spp.; Syncytium of Globodera, Heterodera spp. and Rotylenchulus reniformis and Nurse cells of Tylenchulus semipenetrans) (Mitchum et al., 2012).

1.5 Cyst Nematodes

Cyst nematodes include the genera Heterodera and Globodera. Nematodes belonging to this group are devastating pests of various crops of great economic importance worldwide. Most cyst nematodes tend to have a narrow host range compared to RKN species (Abad and Williamson, 2010). The crops which most impacted by cyst nematodes are potato, which is attacked by the potato cyst nematodes (PCN), Globodera rostochiensis and G. pallida. On an average 9% losses in total potato production are attributed to PCN (Turner and Rowe, 2006). Sugar beet which is attacked by the beet cyst nematode (BCN), H. schachtii; wheat which is attacked by the cereal cyst nematodes (CCN) H. avenae, H. latipons and H. filipjevi. Another major crop for which cyst nematodes cause significant yield losses is soybean due to the soybean cyst nematode (SCN), H. glycines (Jones et al., 2013). Annual losses caused by SCN in USA alone are in excess of $US 1.5 billion (Chen et al., 2001).

The disadvantage of cyst nematodes having a narrow host range is compensated by extreme survival abilities which allow the nematode to persist in the absence of a host for a prolonged period. The cyst itself is formed by the body wall of the dead female and contains the eggs inside. The cyst is usually pale to dark brown in colour due to tanning of the female cuticle

Chapter 1: Introduction

and is usually found either attached to host roots or in the soil. Eggs within cysts can survive for many years in the soil. For example, PCN can survive in the soil in a dormant condition in the absence of a host for 20 years (Eves-van den Akker et al., 2016; Lilley et al., 2005). This degree of survival makes them difficult to manage with crop rotation. Use of natural resistance is the most cost effective and environmentally friendly method of controlling cyst nematodes but identifying and incorporating resistance into many crops is a challenge. In addition evolution of virulent populations (pathotypes) which break resistance presents further problems (Lilley et al., 2005).

1.6 General life cycle of cyst nematodes

The life cycle of PPNs consists of 4 juvenile stages and an adult stage (male or female). The first stage juvenile (J1) develops from the embryo inside the egg. The J1 moults inside the eggshell to the second stage pre-parasitic juvenile (J2) (Curtis, 2007). The life cycle resumes with hatching of the J2s from the eggs within the cyst (Figure 1.5). Most cyst nematode J2s hatch in response to host stimuli in the form of root exudates. In the absence of a host plant, these unhatched J2s enter a diapause stage which remains dormant until the onset of favourable conditions. For some cyst nematodes that develop in situations where a host is likely to be present at all times, this dormancy may be lost. Upon hatching, J2s move in search of the host plant root under the influence of host root stimuli. J2s have lipid reserves in the body which are used as source of energy during the host finding process (Robinson et al., 1987). However, J2s cannot survive for long in the soil. During this host finding process, a pair of amphids in the nematode head region plays a crucial role as chemoreceptors. A range of phytohormones such as auxin and cytokinin, which are found in host root exudates, trigger changes in the surface coat of J2s. It is hypothesized that these changes in cuticle alter the nematode behaviour and prepare them for root invasion (Akhkha et al., 2002; Curtis, 2007). Once it locates the host root, the J2s penetrate the root tissue by mechanical stylet thrusting together with release of enzymes from the gland cells which soften the cell wall. J2s penetrate just behind root tip, in the elongation zone.

Chapter 1: Introduction

Figure 1.5: Typical life cycle of an amphimictic cyst nematode H. schachtii. (Source: Grundler, F.M.W.).

Parasitic J2s migrate intracellularly through the cortical tissues causing significant damage to the root. Once inside the vascular cylinder, the nematode establishes its feeding site, called a syncytium, on which it continues to feed until the completion of its life cycle. The nematode goes through three additional moults to develop into either the adult male or female. Sex is determined by epigenetic factors including environmental conditions and food availability. As feeding progresses the female’s syncytia enlarges, and the females start to swell and become saccate. These white females continue to grow and break out through the root surface, exposing the posterior part of the body outside the root. After successful initiation of parasitism, the nematodes which are destined to be males, feed only to the J3 stage after th which they undergo two subsequent moults in the cuticle of the J3 stage. After the 4 moult, the males revert to the vermiform body shape and emerge from the moulted cuticle in search of females and start mating with females which remain sedentary. Most cyst nematodes reproduce sexually except for H. sacchari, which reproduces by mitotic parthenogenesis (Anonymous, 2014). Shortly after fertilising the females, the male nematodes will die as they are not able to feed again on the host root while the female continues to feed from the syncytium. Just after mating, the fully developed female starts to deposit eggs inside her body. The number of eggs deposited varies from 200 to 500 depending upon the nutrients obtained and other stresses. In a few cyst nematode species, eggs are also deposited outside the body in a gelatinous matrix (e.g. pigeon pea cyst nematode, H. cajani (Gaur et al., 1996)). Nematodes that exhibit such behaviour tend to either have a broad host range or live in an environment in which a host is always available and thus have a reduced requirement for a long-term survival strategy. After the eggs are deposited, and as females start to mature, their

Chapter 1: Introduction

colour changes from white to dark brown due to tanning controlled by the action of polyphenol oxidase enzymes. The female subsequently dies and forms the cyst, which detaches from the roots and drops into the soil. Most cyst nematodes complete their life cycle in about 6 to 8 weeks, although this varies depending on environmental conditions. Some of the eggs start hatching immediately if a host crop is available giving the nematode a chance to complete multiple infection cycles during a host crop season, while some eggs remain dormant for the next cropping seasons (Lilley et al., 2005).

1.7 Syncytia morphology and physiology

Successful parasitism depends on whether the interaction between host and nematode is compatible or incompatible. If the interaction is compatible, the nematode is able to parasitize the plant successfully and complete its life cycle. But if the interaction is incompatible, the nematode fails to establish parasitism and will eventually die without completing its life cycle. This incompatible interaction may be because of a resistant host reaction or may be the result of an attempt to infect a non-host crop. The feeding structure of cyst nematodes, the syncytium, is formed from plant root cells which are modified by the nematode. The syncytium is formed in the inner cortical cells (Globodera spp.) or in the vascular cylinder (Heterodera spp.) (Wyss, 1992). Once inside the cortical tissues/vascular cylinder, the behaviour of J2s changes. During migration the J2s will migrate destructively, piercing cells on its path with the stylet. However, once the nematode is in the cortical tissues or vascular cylinder it starts probing individual cells and waits for the cell response. J2s will retract the stylet if the protoplast of the punctured cell starts collapsing or if deposition of callose-like material occurs near the stylet (Wyss, 1992; Sobczak et al., 1999). This process is repeated until the J2 finds a suitable cell which does not respond adversely to its probing. This initial feeding cell is called the initial syncytial cell (ISC) (Wyss and Zunke, 1986; Wyss, 1992; Golinowski et al., 1997).

Once the ISC is selected, the nematode becomes sedentary and is thought to secrete a cocktail of proteins from its esophageal/ pharyngeal glands cells into the ISC through the stylet aperture (Vanholme et al., 2004). Although the nematode stylet does not appear to penetrate the plasma membrane of the ISC, it is clear that the nematode can deliver these effector proteins into the apoplast and into the cytoplasm of ISC (Mitchum et al., 2013). The ISC starts to increase in size under the influence of the nematode secretions. Localized dissolution of the ISC wall occurs at the plasmodesmata and subsequently fusion of protoplast of

Chapter 1: Introduction

adjoining cells occurs (Grundler et al., 1998; Goverse and Smant, 2014). This process continues until several hundred cells become fused together, giving rise to one continuous symplast (Jones, 1981). Other changes, including enlargement of the nucleus and nucleolus of ISC occur due to multiple rounds of DNA endoreduplication in the absence of nuclear division (Wyss, 1992; Niebel et al., 1996; Goverse et al., 2000). The cytoplasm of the syncytium becomes dense and metabolically active, the large central vacuole breaks down into many small vesicles along with proliferation of smooth endoplasmic reticulum, ribosomes, an increase in the number of mitochondria and plastids (Sobczak et al., 1999; Lilley et al., 1997; Gheysen and Fenoll, 2002). The syncytium also incorporates some conductive tissues which help in efficient transport of water and nutrients to the syncytia (Figure 1.6) (Sobczak and Golinowski, 2011).

Figure 1.6: Syncytium formed by the soybean cyst nematode, Heterodera glycines in soybean root (N- Nematode, S-Syncytium) (Source- American phytopathological society).

In general, the syncytium associated with male nematodes is smaller than those induced by females. Nematode sex determination is governed by various epigenetic factors including environmental conditions and quality and amount of food available to the parasitic nematode (Grundler et al., 1991; Betka, 1991). Each nematode juvenile (J2) can induce only one syncytium (Haegeman et al., 2012). Therefore, formation and maintenance of this nutritive sink is very important for cyst nematodes as it represents the sole source of nourishment for the nematode (Lee et al., 2011). The syncytium will be maintained for as long as the nematodes are attached to it and actively feeding. The syncytium starts degenerating once the nematode stops feeding on it, suggesting involvement of a constant stimulus from the nematode for its maintenance (Sobczak and Golinowski, 2011). When feeding, cyst nematodes produce a feeding tube which connects the nematode stylet opening to the cytoplasm of the syncytium. The nature, formation and role of the feeding tube is not

Chapter 1: Introduction

completely understood but it is believed that it functions as molecular sieve and helps prevent blockage of stylet with subcellular organelles (Davis et al., 2004; Böckenhoff and Grundler, 1994). However, given that the syncytium needs to be kept alive for the duration of the nematode life cycle, the feeding tube may also function to prevent the nematode from destroying the syncytium during feeding.

1.8 Nematode effectors

Plant pathogens including bacteria, fungi, oomycetes and nematodes secrete various proteins, and small molecules into the host plant which aid in infection and other processes essential for successful parasitism (Hogenhout et al., 2009). These secretions are called effectors. Hogenhout et al. (2009), define effectors as all pathogen/ pest proteins and small molecules that alter host-cell structure and function. In the case of nematodes, effectors are the secretions from nematodes which cause changes in the plant system leading to formation of the syncytium or which may activate or suppress defence responses. In most studies on PPN, the term effectors or “parasitism genes” is used to denote nematode secretions which facilitate the compatible interaction with its host plant (Mei et al., 2015; Davis et al., 2008; Rosso et al., 2011). It should be noted that not all proteins secreted by the nematode into the host plant are effectors. Effectors can be identified using a combination of bioinformatics and lab studies. When analysing data for the presence of candidate effectors, two important features are assessed. Firstly, the predicted protein should have an N terminal signal peptide and secondly it should lack a transmembrane domain. The presence of an N terminal signal peptide suggests that the protein is secreted from the cell through the classical secretory pathway via the ER and Golgi. Secreted proteins can have two fates – they may become extracellular or may become embedded in the membrane of the cell. Analysing for the absence of a transmembrane domain shows that the predicted protein is not a membrane protein and may therefore be an extracellular protein, a requirement of an effector. Secreted proteins can be confirmed as effectors by localising the transcript in the secretory organs of the nematode by in situ hybridization (ISH). If the expression of the gene increases during parasitic stages of nematode this indicates a potential role in parasitism, another key feature of an effector.

Effectors can originate from a variety of nematode tissues including the hypodermis and the amphids. However, the major source of nematode effectors is the pharyngeal gland cells,

Chapter 1: Introduction

which produce effectors that are introduced into the host via the stylet (Figure 1.7) (Haegeman et al., 2012).

(a) (b)

Figure 1.7: Illustration of Anterior part of juvenile and developing female. (a) Anterior part of juvenile illustrating single dorsal and two subventral esophageal gland cells that produce the majority of nematode effectors. (b) developing female nematode with enlarged dorsal gland cell and reduced subventral gland cells (Davis et al., 2004).

These effector molecules hijack the plant cell’s metabolic processes and re-programme these for its own benefit. Effectors play a pivotal role in nematode pathogenesis in the following processes: helping J2s during penetration and migration inside the host root tissue, induction and maintenance of feeding site and suppression of host defence response (Curtis, 2007; Gheysen and Mitchum, 2011; Smant and Jones, 2011).

Three specialised glands in the nematode esophagus acts as the principle source of effectors (Hussey, 1989; Davis et al., 2008; Rosso et al., 2011). Each gland is a single celled structure, with a long cytoplasmic extension which terminates anteriorly into an ampulla. This ampulla serves as reservoir of granular secretions and is connected to the esophageal lumen. The gland secretions are released outside the nematode body through the stylet by the action of the metacorpus pump (Hussey, 1989). Distinct morphological changes in the gland cells during nematode parasitism reflect their different functional roles. The two subventral gland cells are highly active during the early stage of infection i.e. assisting nematodes during penetration, migration and initiation of feeding cells. By contrast, the lone dorsal gland is active just after migration ceases, and probably plays a key role in producing effectors that are required for subsequent development and maintenance of the feeding structure. However, the functional boundary between these gland cell types is not clearly demarcated (Mitchum et al., 2013; Haegeman et al., 2012; Davis et al., 2008; Hussey and Mims, 1990).

Chapter 1: Introduction

1.9 Identification of nematode effectors

Effectors play a vital role in nematode parasitism. Therefore, identification of various candidate genes encoding nematode effector proteins is crucial to decipher the plant- nematode interaction. However, this is difficult due to the microscopic size of nematodes, their obligate parasitism, long life cycle and difficulty in performing genetic analysis (Rehman et al., 2016). Their obligate parasitic nature makes it difficult to produce parasitic stages in large numbers and being inside the root makes them difficult to handle for various analyses (Curtis, 2007).

Several techniques have been used for identification of nematode effector proteins. The cDNA-AFLP technique was used to identify and clone three putative effectors namely A4, A18, and A41 from PCN, G. rostochiensis (Qin et al., 2000). The first report of an endogenous animal cellulose gene (beta-1, 4-endoglucanase) from two cyst nematodes, G. rostochiensis and H. glycines was an effector that was identified using monoclonal antibodies (Smant et al., 1998); while polyclonal antibodies were used to identify three genes of the SPRYSEC family (SPRYSEC-9, 15 and 18) from G. rostochiensis (Rehman et al., 2009). Elling et al. (2009), used Expressed Sequence Tags (ESTs) in combination with microarray expression data of different life stages from H. glycines to identify 788 genes upregulated upon infection including several candidate effectors (Elling et al., 2009). ESTs available from different life stages of nematode (Jones et al., 2009) or from aspirated gland cells (Hussey et al., 2011) have been used extensively to identify nematode effectors. Up until February 2015, ~192,230 ESTs were available in sequence datasets from sedentary and migratory parasitic nematodes (Rehman et al., 2016). Several effectors including pectate lyase, expansins, chorismate mutase, polygalacturonase etc. were identified using nematode ESTs (Jones et al., 2009).

During the last decades a lot of investment has been made in genome and transcriptome analysis of PPNs (Rosso et al., 2009). Many PPN transcriptomes have been published including Nacobbus aberrans (Eves-van den Akker et al., 2014); Heterodera avenae (Kumar et al., 2014; Yang et al., 2017); H. schachtii (Fosu-Nyarko et al., 2016); Globodera pallida (Cotton et al., 2014); G. rostochiensis (Eves-van den Akker et al., 2016); Rotylenchulus reniformis (Eves- van den Akker et al., 2016); Meloidogyne graminicola (Haegeman et al., 2013); M. incognita (Danchin et al., 2013); Hirschmanniella oryzae (Bauters et al., 2014) and Pratylenchus penetrans (Vieira et al., 2015). Recently, Maier et al. (2013), developed a

Chapter 1: Introduction

technique for isolation of individual esophageal gland cells from PPNs. They isolated esophageal glands from 9 different PPNs for transcriptome analysis (Maier et al., 2013). Following on from transcriptome projects, several species of PPN have now been the subject of genome sequencing projects. Meloidogyne incognita was the first PPN for which the entire genome was sequenced (Abad et al., 2008). In the same year the genome of another RKN, M. hapla, was also published (Opperman et al., 2008). Genomes of Bursaphelenchus xylophilus (Kikuchi et al., 2011); M. floridensis (Lunt et al., 2014); G. pallida (Cotton et al., 2014); Pratylenchus coffeae (Burke et al., 2015); G. rostochiensis ( Eves- van den Akker et al., 2016); Ditylenchus destructor (Zheng et al., 2016); G. ellingtonae (Phillips et al., 2017); M. arenaria, M. enterolobii and M. haplanaria (Szitenberg et al., 2017) were published and several others are in progress. These transcriptome and genome sequences have accelerated the process of identification of candidate effectors. These studies have also thrown light on the evolution of nematode genomes, in terms of which parts are conserved and which parts have evolved for parasitism.

1.10 Functional validation of effectors

During the last decade, many genes have been identified as putative nematode effectors and the list of candidates is expanding as further genome and transcriptome projects are completed. However, most of the candidate effector genes are novel as there is no match with functionally annotated proteins in the database, making an assessment of their likely function difficult. Various techniques are currently being used for validating the roles of nematode effectors.

RNA interference (RNAi) is one of the most frequently used techniques for gene characterization in C. elegans. It is a widely accepted tool for functional validation due to its high specificity. RNAi allows expression of the targeted transcript gene to be knocked out or reduced and the impact of removing this sequence on the biology of the nematode to be determined. This techniques was used with PPN for the first time to determine gene function in two cyst nematodes (H. glycines and G. pallida) (Urwin et al., 2002). In the case of PPN, RNAi can be used in two ways. Urwin et al. (2002), soaked the infective juveniles of H. glycines and G. pallida in a solution containing double-stranded RNA (dsRNA), complementary to the target gene to be silenced. In the same solution they added octopamine to stimulate uptake by J2s as pre-parasitic J2s do not begin feeding until they are within the plants. They found that RNAi treatment was effective in reducing the transcript abundance in

Chapter 1: Introduction

all treatments with a change in sex ratio observed after targeting a cysteine proteinase gene while targeting a C-type lectin resulted in a 59% reduction in parasitism. Targeting the major sperm protein (MSP) gene reduced transcript abundance in males (Urwin et al., 2002). In another method called host induced gene silencing (HIGS), nematodes are grown on plants that produce a short fragment of dsRNA as a hairpin and ingest this dsRNA when feeding. Using this HIGS approach, Yadav et al. (2006), targeted two different genes (splicing factor and integrase) of M. incognita and demonstrated that host derived dsRNA can also work effectively to control RKN (Yadav et al., 2006). Huang et al. (2006), used both in vitro and in vivo approaches to target the 16D10 effector gene of RKN. In the in vitro approach, J2s were soaked in dsRNA solution while in the case of in vivo, transgenic Arabidopsis lines were generated expressing 16D10 dsRNA. Both the approaches successfully silenced 16D10 and gave resistance to four major RKN species (M. incognita, M. javanica, M. arenaria and M. hapla) (Huang et al., 2006). Transgenic Arabidopsis thaliana lines expressing dsRNA complementary to 8D05 gene of M. incognita resulted in a 90% reduction in root knot infection, which confirms that Mi8D05 gene is crucial for parasitism success of M. incognita (Xue et al., 2013).

In planta localization of nematode effectors by immunolocalization can contribute to functional studies. Immunolocalization studies can be done on nematode infected roots using specific antibodies against the nematode protein. Wang et al. (2010), produced specific antibodies against H. glycines CLEs (HgCLE) peptides and immunolocalized these in the H. glycines infected roots (Wang et al., 2010). A similar approach was used to show that an effector of M. incognita targets the nucleus of the host (Jaouannet et al., 2012).

Over expression of candidate effectors in plants may also provide a route to analyse function. For example, overexpression of the 10A07 gene of H. schachtii in A. thaliana results in higher infection levels, indicating importance of this gene in the host-nematode interaction (Hewezi et al., 2015). In cases where nematode effectors act as a mimic of a plant protein, the function of the effector can be analysed by investigating whether the nematode gene can partially or fully rescue the plant phenotype by complementing the function of the sequence in a mutant line. For example, Arabidopsis mutants for a CLE gene (CLV3) were partially or fully rescued by expressing H. glycines CLE gene (Hg-SYV46) (Wang et al., 2005).

Yeast two hybrid (Y2H) screening is a well-established system used to detect physical interactions between the proteins and is widely used to identify host target proteins of

Chapter 1: Introduction

effectors. Y2H relies on the expression of a reporter gene which is activated by binding of a specific transcription factor. The transcription factor consists of a DNA binding domain (BD) and the activation domain (AD). The protein under investigation is fused to the BD, which acts as bait while a protein/cDNA library is fused to the AD and serves as prey. Expression of the reporter gene will be activated when both the BD and AD of the transcription factor are present in the upstream region of the reporter, which is possible only when bait and prey proteins interact with each other. Using the Y2H approach Hewezi et al. (2008), identified pectin methylesterase 3 (PME3; At3g14310) from A. thaliana as an interactor of the cellulose binding protein (CBP) from H. schachtii (Hewezi et al., 2008). Similarly, the H. schachtii 10A06 effector protein was identified as interacting with Spermidine Synthase2 (SPDS2) from A. thaliana (Hewezi et al., 2010).

1.11 Classification of nematode effectors

Effectors can be categorised into three different types based on the function they perform in planta, as cell wall modifying, defence modulating and development altering effectors.

Cell wall modifying effectors

The cell wall modifying effectors are one of the largest groups of effectors identified from PPNs. This group of enzymes is not usually found in animals and is therefore believed to have been acquired from various prokaryotes through multiple horizontal gene transfer (HGT) events. The plant cell wall is made up of various polysaccharides including cellulose, hemicellulose, pectin, lignin and various other proteins which serve as a protective shield against any potential intruder. In order to penetrate and migrate inside the host root, pre- parasitic juveniles need to cross the barrier of the cell wall. Along with mechanical stylet thrusting, the nematode secretes various enzymes from its esophageal glands which modify cell wall by making it softer or degrading glycosidic bonds.

The first effector protein identified from PPN was a beta-1,4-endoglucanase which degrades the cellulose component of plant cell wall (Smant et al., 1998). Several cellulase family proteins especially (mostly from glycosyl hydrolase family 5 (GHF5)) have been identified from other PPN genera including Globodera, Heterodera, Ditylenchus, Radopholus, and Meloidogyne (Abad et al., 2008). Other than cellulases, several other effector proteins have been identified that act on the host cell wall including pectate lyase which acts on pectate sugar polymers; polygalacturonases that degrade galacturonans; xylanases which hydrolyse

Chapter 1: Introduction

xylans; Arabinogalactan endo-1,4-beta- galactosidase and arabinose which hydrolyse polysaccharides from pectin sidechains.

Cellulose binding and expansin like proteins have been reported in sedentary endoparasites (Ding et al., 1998; Gao et al., 2004; Wieczorek et al., 2006). Both of these proteins were found to weaken the host cell wall and these interactions may accentuate the activity of other cell wall degrading enzymes (Hassan et al., 2010). In addition, one CBP effector from H. schachtii (Hs-CBP) interacts with a plant Pectin Methylesterase Protein3 (PME3). Overexpression of Hs-CBP in A. thaliana was found to increase PME3 activity in planta, which in turn reduces level of methylesterified pectin in cell walls enabling the access of other cell wall modifying/degrading enzymes to cell wall polymers during feeding cell formation (Hewezi et al., 2008). This shows that cell wall modifying proteins can have a range of roles in plant-nematode interactions.

Defence modulating effectors

Unlike mammals, plants do not have mobile defender cells. Instead each cell has innate immunity and is able to mount a defence response against intruders. Plants have evolved a multi-layered immune system to ensure pathogen recognition and activation of defences. In the first layer of defence, plants deploy various transmembrane pattern recognition receptors (PRRs) to survey the apoplast for the presence of pathogens. PRRs recognise conserved microbial/pathogen-associated molecular patterns (MAMPs/PAMPs) from pathogens or breakdown products of the cell wall generated by pathogens (damage associated molecular patterns – DAMPs) and activate PAMP triggered immunity (PTI). PTI is a general response to any pathogen that aims to curb pathogen growth and promote disease resistance (Jones and Dangl, 2006). PTI leads to cell wall thickening by lignification and callose deposition along with production of reactive oxygen species at the site of infection (Hewezi and Baum, 2012; Holbein et al., 2016). To tackle PTI, virulent/ evolved pathogens secrete effector molecules which suppress this response thereby leading to effector-triggered susceptibility (ETS) (Mantelin et al., 2017). The second layer of defence is activated when pathogen effectors or their activity are recognised by resistance proteins from the plant and evoke effector triggered immunity (ETI). ETI invokes a strong localised cell death at the site of infection known as hypersensitive response (HR) (Jones and Dangl, 2006; Mei et al., 2015). Sedentary PPNs need to subvert plant defences in order to protect themselves as well as their feeding

Chapter 1: Introduction

structures. During last few years, many nematode effectors involved in defence suppression were identified.

G. rostochiensis produces glutathione peroxidase and peroxidoxin which metabolise hydrogen peroxide, a plant defence compound produced as result of the reactive oxygen burst (Robertson et al., 2000; Jones et al., 2004). Venom allergen-like protein from the same nematode (Gr-VAP1) bind to and inhibit Rcr3pim, a papain-like cysteine protease, resulting in an increase in susceptibility in tomato plants (Lozano-Torres et al., 2014). Another G. rostochiensis effector from the SPRYSEC family supresses plant defence response (Postma et al., 2012; Rehman et al., 2016). Subsequent studies showed that several other SPRYSEC effectors can suppress host defences (Mei et al., 2015). Ali et al. (2015), identified 2 effectors from G. rostochiensis namely, ubiquitin extension protein (GrUBCEP12) and an expansin- like protein (GrEXPB2). These both effectors were found to suppress plant cell death responses initiated in plant defences when transiently expressed in tobacco (Ali et al., 2015). The fatty acid and retinol binding protein-1 from G. pallida (Gp-FAR-1) and of M. javanica (Mj-FAR-1), present on surface coat of nematodes are involved in disrupting the jasmonic acid (JA) signalling pathway (Prior et al., 2001; Iberkleid et al., 2013). Both SA and JA play key role in plant defence signalling (Haegeman et al., 2012).

One H. schachtii effector, 10A06 was found to interact with spermidine synthase (SPDS2) in planta. This interaction results in a change in polyamine synthesis and increase in spermidine content and subsequent polyamine oxidase (PAO) activity, thereby reducing ability of hosts to produce defence related compounds such as salicylic acid. It has also been found that constitutive expression of the H. schachtii 10A06 in Arabidopsis thaliana significantly increases susceptibility to nematodes as well as to unrelated pathogens including Pseudomonas syringae pv tomato (Pst DC300) and the yellow strain of cucumber mosaic virus (Hewezi et al., 2010). Another effector from H. schachtii, Hs-30C02 was identified as homologue of soybean cyst nematode, H. glycines gene Hg-30C02. This gene specifically interacts with host AT4G16260 gene and interferes in its role as a plant pathogenesis related protein (PR-2). It also suppresses β-1, 3-endoglucanase activity which in turn increases plant susceptibility to the nematode. Arabidopsis mutants for the AT4G16260 gene showed high susceptibility while lines overexpressing same gene were found to be resistant to H. schachtii (Hamamouch et al., 2012). A homologue of the H. glycines 4F01 effector identified from H. schachtii (Hs4F01) was found to interact with an Arabidopsis oxidoreductase member of the

Chapter 1: Introduction

2OG-Fe(II) oxygenase family and promote susceptibility to nematodes and oomycetes (Patel et al., 2010).

Cyst nematodes, RKN and P. coffeae have been identified to produce chorismate mutase (CM), a key enzyme in the shikimate pathway in plants. The chorismate mutase from nematode may act on plant chorismate and convert it to prephenate. This may lead to reduced availability of chorismate for conversion to salicylic acid (SA) and thus may prevent triggering of host defence response (Doyle and Lambert, 2003).

Development altering effectors

As described above, sedentary PPN inject effectors into the host which profoundly modify host cell structure and physiology. Nematode effector proteins emulating plant CLAVATA3/ENDOSPERM SURROUNDING REGION-related (CLE) proteins have been reported from G. rostochiensis and H. schachtii (Lu et al., 2009; Wang et al., 2005). One CLE peptide from G. rostochiensis (GrCLE1) which mimics plant CLE, is processed into its active form by host plant proteases. GrCLE1 binds with plant CLAVATA2-like receptor (StCLV2) from potato and alters the plant root phenotype (Guo et al., 2011; Chen et al., 2015). A nematode CLE has been shown to complement an Arabidopsis mutant lacking this protein showing that the nematode sequences are functional CLE proteins (Wang et al., 2005).

Another effector has been identified that may interact with hormone signalling in plants. The orthologue of soybean cyst nematode H. glycines 19C07 effector has been characterised from H. schachtii (Hs19C07) as an interactor of the Arabidopsis auxin influx transporter LAX3. This interaction may allow the nematode to modulate the auxin influx inside the roots, thus stimulating cell wall hydrolysis for development of syncytium (Lee et al., 2011). Similarly, the in planta expression of chorismate mutase from M. javanica in hairy roots of soybean affects formation and development of the root vascular system, possibly by affecting auxin levels in the host (Doyle and Lambert, 2003).

Nematodes may also themselves produce plant hormones in order to manipulate their hosts. Siddique et al. (2015), identified an isopentenyl transferase gene from H. schachtii that may be involved in synthesis of cytokinin, suggesting that juveniles of H. schachtii can produce functional cytokinin. They showed that cytokinin is important for development of the syncytia and that silencing of this gene results in reduction in infection and as well as improper syncytia development (Siddique et al., 2015).

Chapter 1: Introduction

1.12 Comparative genomics

Each nematode genome consists of set of genes required for essential life functions and others which are more diverged to define unique biology of each species (Blaxter et al., 2011). Comparing genomes between different genera of plant parasites allows genes conserved across nematode genera that may be essential for parasitism, including core effectors, to be identified. Comparisons between closely related species that have different host specificities may produce information about the genes governing host range. Genome comparisons can also be analysed to understand horizontally transferred effector genes, expansion or reduction of gene families, evolution of new effectors and evolution of pathotypes.

Recent advances in high-throughput sequencing coupled with the reduced costs associated with this, has resulted in increased numbers of nematode genomes and transcriptomes being analysed. In addition, tools for analysis of genomes and transcriptomes have been developed and are increasingly accessible to non-specialists. Up till now (2017), eight species of PPNs (M. incognita, M. hapla, M. floridensis, G. pallida, G. rostochiensis, B. xylophilus, D. destructor and P. coffeae) had completed genome sequences and more than 30 other sequencing projects are in progress (Zheng et al., 2016; Kikuchi et al., 2017).

Phylogenetic analysis has shown that plant parasitism by nematodes has evolved at least four times independently. PPNs are found in four (Clade I, II, 10b and 12) out of twelve clades defined by Van Megen et al. (Van Megen et al., 2009). Compared to free living counterparts (C. elegans genome size- 100 Mb), parasitic nematodes (APNs & PPNs) contain reduced numbers of genes, reflecting a dependency on the host for compounds required for development. So far, the highest level of genome reduction has been observed in root lesion nematode P. coffeae which is reported to have a genome size of 19.7 Mb (Burke et al., 2015). Similarly, sedentary endoparasites (RKN and cyst nematode), which spend most of their life cycle inside their host roots, have a reduced number of genes associated with immunity against pathogens, possibly reflecting the fact that they are protected from other pathogens by being within their hosts (Kikuchi et al., 2017).

To be successful plant parasites, in addition to their housekeeping genes, nematodes need to have new genes for this new function. These new genes can be acquired from other organisms through HGT event or by modifying housekeeping genes through duplication followed by diversification. In addition, entirely new sequences may evolve with specific

Chapter 1: Introduction

roles in plant parasitism. Several genes have been acquired by nematodes from bacteria and fungi through the process of HGT. PPNs from all three clades studied to date have acquired cell wall degrading proteins (CWDP) but each clade has different CWDP suggesting multiple HGT events in nematodes (Jones et al., 2005).

Housekeeping genes are required for basic cellular function. A modified form of ubiquitin, an important housekeeping gene found in all eukaryotes has been identified from gland secretions of several species of cyst nematodes. One such protein from G. rostochiensis, GrCEP12 is able to supress the flg22 induced PTI response, while its over-expression increases susceptibility to nematode and Streptomyces scabies, an unrelated bacterial pathogen in potato, suggesting a role in defence suppression (Chen et al., 2013). Similarly, SPRY domain proteins, common in eukaryotes, occur with other functional protein domains and are adapted to function as effectors in cyst nematodes. The SPRYSEC gene family is highly expanded in PCNs (Mei et al., 2015). Several SPRYSEC proteins were identified as defence suppressors (Rehman et al., 2009; Diaz-Granados et al., 2016).

The interactions between hosts and parasites are under strong selection pressure. Possibly reflecting these selection pressures many effectors evolve de novo in PPNs (and other plant pathogens). This is reflected by the fact that, other than cell wall degrading enzymes, there is almost no overlap between effectors of cyst nematodes and RKN (Thorpe et al., 2014).

1.13 Aims of the thesis

In order to gain more knowledge about the core effectors conserved across cyst nematodes, we plan to compare the effectors present in three different species of cyst nematode: Heterodera sacchari - a parasite of rice on one side, with H. schachtii - a parasite of sugar beet and G. pallida – a parasite of potato on other side. Our specific interest will be the comparison of cyst nematodes that infect monocots and dicots. A lot of information regarding the effector repertoires of H. schachtii (Fosu-Nyarko et al., 2016) and G. pallida (Cotton et al., 2014; Thorpe et al., 2014) is already available.

H. sacchari is commonly known as the sugarcane cyst nematode as it was reported for the first time as a parasite of sugarcane crop, Saccharum officinale L., in Congo-Brazzaville. Later, in 1970, the same nematode was found to infect rice (Oryza sativa). H. sacchari is one of the most important parasites and limiting factors of sugarcane and rice production. Apart from sugarcane and rice, it also infects and multiplies on several weed hosts of the poaceae / gramineae family. H. sacchari has a wide distribution and has been reported from Africa

Chapter 1: Introduction

(Benin, Burkina Faso, Chad, Congo, Côte d’Ivoire, Ghana, Liberia, Nigeria, Senegal and The Gambia), from Asia (India, Pakistan and Thailand) and from Jamaica (Anonymous, 2014; Ferris, 2005). Although H. sacchari is reported under both upland and lowland rice cultivations it has a greater impact in upland conditions (Coyne and Plowright, 2000; Bridge et al., 2005).

To date no studies have been done on the molecular interaction between H. sacchari and rice. We have therefore planned our research with following objectives:

1. To identify effectors from the sugarcane cyst nematode H. sacchari and compare those present in this nematode to those from the dicot pathogens.

2. To analyse conservation of function of effectors of H. sacchari compared to those from H. schachtii and G. pallida.

3. To analyse the potential role of changes in effectors function in determining host range.

1.14 References

Abad, P., Gouzy, J., Aury, J.-M., Castagnone-Sereno, P., Danchin, E.G.J., Deleury, E., Perfus-Barbeoch, L., Anthouard, V., Artiguenave, F., Blok, V.C., Caillaud, M., Coutinho, P.M., Dasilva, C., De Luca, F., Deau, F., Esquibet, M., Flutre, T., Goldstone, J.V., Hamamouch, N., Hewezi, T., Jaillon, O., Jubin, C., Leonetti, P., Magliano, M., Maier, T.R., Markov, G.V., McVeigh, P., Pesole, G., Poulain, J., Robinson-Rechavi, M., Sallet, E., S'egurens, B., Steinbach, D., Tytgat, T., Ugarte, E., Ghelder, C.V., Veronico, P., Baum, T.J., Blaxter, M., Bleve-Zacheo, T., Davis, E.L., Ewbank, J.J., Favery, B., Grenier, E., Henrissat, B., Jones, J.T., Laudet, V., Maule, A.G., Quesneville, H., Rosso, M.N., Schiex, T., Smant, G., Weissenbach, J. and Wincker, P. (2008). Genome sequence of the metazoan plant-parasitic nematode Meloidogyne incognita. Nature Biotechnology 26, 909–915.

Abad, P. and Williamson, V.M. (2010). Plant Nematode Interaction: A sophisticated dialogue. In Advances in botanical research (Kader, J.-C. and Delseny, M. eds.), 53 pp. 147–192, Academic Press.

Akhkha, A., Kusel, J., Kennedy, M. and Curtis, R. (2002). Effects of phytohormones on the surfaces of plant-parasitic nematodes. Parasitology 125, 165–175.

Chapter 1: Introduction

Ali, S., Magne, M., Chen, S., Côté, O., Stare, B.G., Obradovic, N., Jamshaid, L., Wang, X., Bélair, G. and Moffett, P. (2015). Analysis of putative apoplastic effectors from the nematode, Globodera rostochiensis, and identification of an expansin-like protein that can induce and suppress host defenses. PLoS One 10, e0115042.

Anonymous (2014). CABI’s report on sugarcane cyst nematode, Heterodera sacchari. (http://download.ceris.purdue.edu/file/3047).

Ayoub, S.M. (1980). Plant nematology: an agricultural training aid. (Revised)., Sacramento, California: NemaAid Publication. pp 195.

Bartlem, D.G., Jones, M.G.K. and Hammes, U.Z. (2014). Vascularization and nutrient delivery at root-knot nematode feeding sites in host roots. Journal of experimental Botany 65, 1789–1798.

Betka, M. (1991). Influence of changes in the nurse cell system (syncytium) on the development of the cyst nematode Heterodera schachtii: single amino acids. Phytopathology 81(1), 75-79.

Blaxter, M. (2011). Nematodes: The worm and its relatives. PLoS Biology 9(4), e1001050.

Blaxter, M., Kumar, S., Kaur, G., Koutsovoulos, G. and Elsworth, B. (2012). Genomics and transcriptomics across the diversity of the Nematoda. Parasite Immunology 34, 108– 120.

Blaxter, M. and Koutsovoulos, G. (2015). The evolution of parasitism in Nematoda. Parasitology 142, S26–S39.

Böckenhoff, A. and Grundler, F.M.W. (1994). Studies on the nutrient uptake by the beet cyst nematode Heterodera schachtii by in situ microinjection of fluorescent probes into the feeding structures in Arabidopsis thaliana. Parasitology 109, 249–255.

Bohlmann, H. (2015). Introductory chapter on the basic biology of cyst nematodes. In Advances in Botanical Research (Escobar, C. and Fenoll, C., eds.) 73, pp. 33–59. Academic Press.

Bridge, J., Plowright, R.A. and Peng, D. (2005). Nematode Parasite of Rice. In Plant Parasitic Nematodes in Subtropical and Tropical Agriculture (Luc, M., Sikora, R.A. and Bridge J, eds.) pp. 87-130.Wallingford, U.K: CAB International.

Chapter 1: Introduction

Burke, M., Scholl, E.H., Bird, D.M.K., Schaff, J.E., Colman, S.D., Crowell, R., Diener, S., Gordon, O., Graham, S., Wang, X., Windham, E., Wright, G.M. and Opperman, C.H. (2015). The plant parasite Pratylenchus coffeae carries a minimal nematode genome. Nematology 17, 621–637.

Chen, S., MacDonald, D.H., Kurle, J.E. and Reynolds, D.A. (2001). The soybean cyst nematode. pdf document of extension service, College of Agricultural, Food, and Environmental Sciences University of Minnesota. pp. 1–8.

Chen, S., Chronis, D. and Wang, X. (2013). The novel GrCEP12 peptide from the plant- parasitic nematode Globodera rostochiensis suppresses flg22-mediated PTI. Plant Signaling and Behavior 8:e25359.

Chen, S., Lang, P., Chronis, D., Zhang, S., Jong, W.S.D., Mitchum, M.G. and Wang, X. (2015). In planta processing and glycosylation of a nematode CLAVATA3/ENDOSPERM SURROUNDING REGION-like effector and its interaction with a host CLAVATA2-like receptor to promote parasitism. Plant Physiology 167, 262–272.

Cotton, J.A., Lilley, C.J., Jones, L.M., Kikuchi, T., Reid, A.J., Thorpe, P., Tsai, I.J., Beasley, H., Blok, V.C., Cock, P.J.A., Eves-van den Akker, S., Holroyd, N., Hunt, M., Mantelin, S., Naghra, H., Pain, A., Palomares-Rius, J.E., Zarowiecki, M., Berriman, M., Jones, J.T. and Urwin, P.E. (2014). The genome and life-stage specific transcriptomes of Globodera pallida elucidate key aspects of plant parasitism by a cyst nematode. Genome Biology 15, R43.

Coyne, D.L. and Plowright, R.A. (2000). Pathogenicity of cyst nematode, Heterodera sacchari, on rice in sand and clay soil. International Rice Research Notes 25, 17–18.

Crompton, D.W.T. (1999). How much human helminthiasis is there in the world? Journal of Parasitology 85, 397–403.

Curtis, R.H.C. (2007). Plant parasitic nematode proteins and the host-parasite interaction. Briefings in Functional Genomics and Proteomics 6, 50–58.

Danchin, E.G.J., Arguel, M.J., Campan-Fournier, A., Perfus-Barbeoch, L., Magliano, M., Rosso, M.N., Rocha, M.D., Silva, C.D., Nottet, N., Labadie, K., Guy, J., Artiguenave, F. and Abad, P. (2013). Identification of novel target genes for safer and more specific

Chapter 1: Introduction

control of root-knot nematodes from a pan-genome mining. PLoS Pathogens 9(10), e1003745.

Davis, E.L., Hussey, R.S. and Baum, T.J. (2004). Getting to the roots of parasitism by nematodes. Trends in Parasitology 20, 134–141.

Davis, E.L., Hussey, R.S., Mitchum, M.G. and Baum, T.J. (2008). Parasitism proteins in nematode-plant interactions. Current Opinion in Plant Biology 11, 360–366.

Decraemer, W. and Hunt, D.J. (2006). Structure and classification In: plant nematology (Perry, R.N., Moens, M. eds.) pp. 3-32. CABI.

Diaz-Granados, A., Petrescu, A.-J., Goverse, A. and Smant, G. (2016). SPRYSEC Effectors: A versatile protein-binding platform to disrupt plant innate immunity. Frontiers in Plant Science 7, 1575.

Ding, X., Shields, J., Allen, R. and Hussey, R.S. (1998). A secretory cellulose-binding protein cDNA cloned from the root-knot nematode (Meloidogyne incognita). Molecular Plant-Microbe Interactions 11, 952–959.

Doyle, E.A. and Lambert, K.N. (2003). Meloidogyne javanica chorismate mutase 1 alters plant cell development. Molecular Plant-Microbe Interactions 16, 123–131.

Elling, A.A., Mitreva, M., Gai, X., Martin, J., Recknor, J., Davis, E.L., Hussey, R.S., Nettleton, D., McCarter, J.P. and Baum, T.J. (2009). Sequence mining and transcript profiling to explore cyst nematode parasitism. BMC Genomics 10, 58.

Elling, A.A. (2013). Major emerging problems with minor Meloidogyne species. Phytopathology 103, 1092–1102.

De Almeida Engler, J., Engler, G. and Gheysen, G. (2011). Unravelling the plant cell cycle in nematode induced feeding sites In: Genomics and Molecular Genetics of Plant- Nematode Interactions (Jones, J., Gheysen, G., Fenoll, C., eds.), pp. 349-368. Springer.

Eves-van den Akker, S., Lilley, C.J., Danchin, E.G.J., Rancurel, C., Cock, P.J.A., Urwin, P.E. and Jones, J.T. (2014). The transcriptome of Nacobbus aberrans reveals insights into the evolution of sedentary endoparasitism in plant-parasitic nematodes. Genome Biology and Evolution 6, 2181–2194.

Chapter 1: Introduction

Eves-van den Akker, S., Lilley, C.J., Jones, J.T. and Urwin, P.E. (2014). Identification and characterisation of a hyper-variable apoplastic effector gene family of the potato cyst nematodes. PLoS Pathogens 10, e1004391.

Eves-van den Akker, S., Lilley, C.J., Yusup, H.B., Jones, J.T. and Urwin, P.E. (2016). Functional C-TERMINALLY ENCODED PEPTIDE (CEP) plant hormone domains evolved de novo in the plant parasite Rotylenchulus reniformis. Molecular Plant Pathology 17, 1265–1275.

Eves-van den Akker, S., Laetsch, D.R., Thorpe, P., Lilley, C.J., Danchin, E.G.J., Rocha, M.D., Rancurel, C., Holroyd, N.E., Cotton, J.A., Szitenberg, A., Grenier, E., Montarry, J., Mimee, B., Duceppe, M., Boyes, I., Marvin, J.M.C., Jones, L.M., Yusup, H.B., Lafond-Lapalme, J., Esquibet, M., Sabeh, M., Rott, M., Overmars, H., Finkers- Tomczak, A., Smant, G., Koutsovoulos, G., Blok V.C., Mantelin, S., Cock, P.J.A., Phillips, W., Henrissat, B., Urwin, P.E., Blaxter M. and Jones, J.T. (2016). The genome of the yellow potato cyst nematode, Globodera rostochiensis, reveals insights into the bases of parasitism and virulence. Genome Biology 1–23.

Ferris (2005). Nemaplex.

Fosu-Nyarko, J., Nicol, P., Naz, F., Gill, R. and Jones, M.G.K. (2016). Analysis of the transcriptome of the infective stage of the beet cyst nematode, H. schachtii. PLoS One 11(1), e0147511.

Gao, B., Allen, R., Davis, E.L., Baum, T.J. and Hussey, R.S. (2004). Molecular characterisation and developmental expression of a cellulose-binding protein gene in the soybean cyst nematode Heterodera glycines. International Journal for Parasitology 34, 1377–1383.

Gaur, H.S., Beane, J. and Perry, R.N. (1996). Effect of storage in vitro and in soil on the hatch from cysts of the pigeonpea cyst nematode, Heterodera cajani. Fundamental and Applied Nematology 19, 537–543.

Gheysen, G. and Fenoll, C. (2002). Gene expression in nematode feeding sites. Annual Review of Phytopathology 40, 191–219.

Gheysen, G. and Mitchum, M.G. (2011). How nematodes manipulate plant development pathways for infection. Current Opinion in Plant Biology 14, 415–421.

Chapter 1: Introduction

Golinowski, W., Sobczak, M., Kurek, W. and Grymaszewska, G. (1997). The structure of syncytia. In: Cellular and Molecular Aspects of Plant–Nematode Interactions (Fenoll, C., Grundler, F.M.W. and Ohl, S., eds.), pp. 80–97. Dordrecht: Kluwer Academic Publishers.

Goverse, A., De Almeida Engler, J., Verhees, J., Krol, S. van der., Helder, J. and Gheysen, G. (2000). Cell cycle activation by plant parasitic nematodes. Plant Molecular Biology 43, 747–761.

Goverse, A. and Smant, G. (2014). The activation and suppression of plant innate immunity by parasitic nematodes. Annual Review of Phytopathology 52, 243–265.

Grundler, F.M.W., Betka, M. and Wyss, U. (1991). Influence of changes in the nurse cell system (syncytium) on sex determination and development of the cyst nematode Heterodera schachtii: total amounts of proteins and amino acids. Phytopathology 81(1), 70-74.

Grundler, F.M.W., Sobczak, M. and Golinowski, W. (1998). Formation of wall openings in root cells of Arabidopsis thaliana following infection by the plant-parasitic nematode Heterodera schachtii. European Journal of Plant Pathology 104, 545–551.

Grundler, F.M.W. and Hofmann, J. (2011). Water and nutrient transport in nematode feeding sites. In: Genomics and Molecular Genetics of Plant-Nematode Interactions. (Jones J., Gheysen G., Fenoll C., eds.), pp. 423-439. Springer, Dordrecht.

Gubanov, N.M. (1951). Giant nematode from the placenta of cetaceans- Placentonema gigantissima nov.gen. nov.sp. Dokl. Akad. Nayuk SSSR 77.

Guerrant, R.L., Walker, D.H. and Weller, P.F. (2011). Tropical infectious diseases: principles, pathogens and practice (Third Edition) pp. 1082–1130. Edinburgh: W.B. Saunders.

Guo, Y., Ni, J., Denver, R., Wang, X. and Clark, S.E. (2011). Mechanisms of molecular mimicry of plant CLE peptide ligands by the parasitic nematode Globodera rostochiensis. Plant Physiology 157, 476–484.

Haegeman, A., Mantelin, S., Jones, J.T. and Gheysen, G. (2012). Functional roles of effectors of plant-parasitic nematodes. Gene 492, 19–31.

Chapter 1: Introduction

Haegeman, A., Bauters, L., Kyndt, T., Rahman, M.M. and Gheysen, G. (2013). Identification of candidate effector genes in the transcriptome of the rice root knot nematode Meloidogyne graminicola. Molecular Plant Pathology 14, 379–390.

Hamamouch, N., Li, C., Hewezi, T., Baum, T.J., Mitchum, M.G., Hussey, R.S., Vodkin, L.O. and Davis, E.L. (2012). The interaction of the novel 30C02 cyst nematode effector protein with a plant β-1,3-endoglucanase may suppress host defence to promote parasitism. Journal of Experimetal Botany 63, 3683–3695.

Hassan, S., Behm, C.A. and Mathesius, U. (2010). Effectors of plant parasitic nematodes that re-program root cell development. Functional Plant Biology 37, 933–942.

Hewezi, T., Howe, P., Maier, T.R., Hussey, R.S., Mitchum, M.G., Davis, E.L. and Baum, T.J. (2008). Cellulose binding protein from the parasitic nematode Heterodera schachtii interacts with arabidopsis pectin methylesterase: cooperative cell wall modification during parasitism. Plant Cell Online 20, 3080–3093.

Hewezi, T., Howe, P.J., Maier, T.R., Hussey, R.S., Mitchum, M.G., Davis, E.L. and Baum, T.J. (2010). Arabidopsis spermidine synthase is targeted by an effector protein of the cyst nematode Heterodera schachtii. Plant Physiology 152, 968–984.

Hewezi, T. and Baum, T. (2012). Manipulation of plant cells by cyst and root-knot nematode effectors. Molecular Plant-Microbe Interactions. 26(1), 9-16.

Hewezi, T., Juvale, P.S., Piya, S., Maier, T.R., Rambani, A., Rice, J.H., Mitchum, M.G., Davis, E.L., Hussey, R.S. and Baum, T.J. (2015). The cyst nematode effector protein 10A07 targets and recruits host posttranslational machinery to mediate its nuclear trafficking and to promote parasitism in Arabidopsis. Plant Cell 27, 891–907.

Hodda M. (2011). "Phylum Nematoda Cobb, 1932. In: Zhang, Z.-Q. (Ed.) Animal biodiversity: An outline of higher-level classification and survey of taxonomic richness". Zootaxa 3148: 63-95.

Hogenhout, S.A., Van der Hoorn, R.A.L., Terauchi, R. and Kamoun, S. (2009). Emerging concepts in effector biology of plant-associated organisms. Molecular Plant-Microbe Interactions 22, 115–122.

Holbein, J., Grundler, F.M.W. and Siddique, S. (2016). Plant basal resistance to nematodes: an update. Journal of Experimental Botany 67, 2049–2061.

Chapter 1: Introduction

Huang, G., Allen, R., Davis, E.L., Baum, T.J. and Hussey, R.S. (2006). Engineering broad root-knot resistance in transgenic plants by RNAi silencing of a conserved and essential root-knot nematode parasitism gene. Proceedings of the National Academy of Sciences, USA 103, 14302–14306.

Huettel, R.N. and Golden, A.M. (1991). Nathan Augustus Cobb: The father of nematology in the United States. Annual Review of Phytopathology 29, 15–27.

Hussey, R.S. (1989). Disease-inducing secretions of plant-parasitic nematodes. Annual Review of Phytopathology 27, 123-141.

Hussey, R.S. and Mims, C.W. (1990). Ultrastructure of esophageal glands and their secretory granules in the root-knot nematode Meloidogyne incognita. Protoplasma 156, 9–18.

Hussey, R.S., Huang, G. and Allen, R. (2011). Microaspiration of esophageal gland cells and cDNA library construction for identifying parasitism genes of plant-parasitic nematodes. In: Plant Immunity-Methods and Protocols. (McDowell J.M. eds.), pp. 89-107, Methods in Molecular Biology book series. Springer publishers.

Iberkleid, I., Vieira, P., de Almeida Engler, J., Firester, K., Spiegel, Y. and Horowitz, S.B. (2013). Fatty acid-and retinol-binding protein, Mj-FAR-1 induces tomato host susceptibility to root-knot nematodes. PLoS One 8(5), e64586.

Jaouannet, M., Perfus Barbeoch, L., Deleury, E., Magliano, M., Engler, G., Vieira, P., Danchin, E.G.J., Rocha, M.D., Coquillard, P., Abad, P. and Rosso, M.N. (2012). A root- knot nematode secreted protein is injected into giant cells and targeted to the nuclei. New Phytologist 194, 924–931.

Jaouannet, M. and Rosso, M.N. (2013). Effectors of root sedentary nematodes target diverse plant cell compartments to manipulate plant functions and promote infection. Plant Signaling and Behavior 8(9), e25507.

Jaubert, S., Milac, A.L., Petrescu, A.J., de Almeida-Engler, J., Abad, P. and Rosso, M.N. (2005). In planta secretion of a calreticulin by migratory and sedentary stages of root- knot nematode. Molecular Plant-Microbe Interactions 18, 1277–1284.

Jones (1981). Host cell responses to endoparasitic nematode attack: structure and function of giant cells and syncytia. Annals of Applied Biology 97, 353–372.

Chapter 1: Introduction

Jones, J.T., Reavy, B., Smant, G. and Prior, A.E. (2004). Glutathione peroxidases of the potato cyst nematode Globodera rostochiensis. Gene 324, 47-54.

Jones, J.T., Furlanetto, C. and Kikuchi T. (2005). Horizontal gene transfer from bacteria and fungi as a driving force in the evolution of plant parasitism in nematodes. Nematology 7, 641–646.

Jones, J.D.G. and Dangl, J.L. (2006). The plant immune system. Nature 444, 323–329.

Jones, J.T., Kumar, A., Pylypenko, L.A., Thirugnanasambandam, A., Castelli, L., Chapman, S., Cock, P.J.A., Grenier, E., Lilley, C.J., Phillips, M.S. and Blok, V.C. (2009). Identification and functional characterization of effectors in expressed sequence tags from various life cycle stages of the potato cyst nematode Globodera pallida. Molecular Plant Pathology 10(6), 815–828.

Jones, J.T., Haegeman, A., Danchin, E.G.J., Gaur, H.S., Helder, J., Jones, M.G.K., Kikuchi, T., Manzanilla-López, R., Palomares-Rius, J.E., Wesemael, W.M.L. and Perry, R.N. (2013). Top 10 plant-parasitic nematodes in molecular plant pathology. Molecular Plant Pathology 14, 946–961.

Kikuchi, T., Cotton, J.A., Dalzell, J.J., Hasegawa, K., Kanzaki, N., McVeigh, P., Takanashi, T., Tsai, I.J., Assefa, S.A., Cock, P.J.A., Otto, T.D., Hunt, M., Reid, A.J., Sanchez- Flores, A., Tsuchihara, K., Yokoi, T., Larsson, M.C., Miwa, J., Maule, A.G., Sahashi, N., Jones, J.T. and Berriman, M. (2011). Genomic insights into the origin of parasitism in the emerging plant pathogen Bursaphelenchus xylophilus. PLoS Pathogens 7(9), e1002219.

Kikuchi, T., Eves-Van Den Akker, S. and Jones, J.T. (2017). Genome evolution of plant- parasitic nematodes. Annual Review of Phytopathology 55, 333–354.

Kiontke, K. and Fitch, D.H.A. (2013). Nematodes. Current biology 23(19), 862-864.

Kumar, M., Gantasala, N.P., Roychowdhury, T., Thakur, P.K., Banakar, P., Shukla, R.N., Jones, M.G.K. and Rao, U. (2014). De novo transcriptome sequencing and analysis of the cereal cyst nematode, Heterodera avenae. PLoS One 9(5), e96311.

Kumar, S., Koutsovoulos, G., Kaur, G. and Blaxter, M. (2012). Toward 959 nematode genomes. Worm 1, 41–49.

Chapter 1: Introduction

Lacey, L.A. and Georgis, R. (2012). Entomopathogenic nematodes for control of insect pests above and below ground with comments on commercial production. Journal of Nematology 44, 218–225.

Lee, C., Chronis, D., Kenning, C., Peret, B., Hewezi, T., Davis, E.L., Baum, T.J., Hussey, R.S., Bennett, M. and Mitchum, M.G. (2011). The novel cyst nematode effector protein 19C07 interacts with the arabidopsis auxin influx transporter LAX3 to control feeding site development. Plant Physiology 155, 866–880.

Lilley, C.J., Urwin, P.E., Atkinson, H.J. and McPherson, M.J. (1997). Characterization of cDNAs encoding serine proteinases from the soybean cyst nematode Heterodera glycines. Molecular and Biochemical Parasitology 89, 195–207.

Lilley, C.J., Atkinson, H.J. and Urwin, P.E. (2005). Molecular aspects of cyst nematodes. Molecular Plant Pathology 6, 577–588.

Lozano-Torres, J.L., Wilbers, R.H.P., Warmerdam, S., Finkers-Tomczak, A., Diaz-Granados, A., Van-Schaik, C.C., Helder, J., Bakker, J., Goverse, A., Schots, A. and Smant, G. (2014). Apoplastic venom allergen-like proteins of cyst nematodes modulate the activation of basal plant innate immunity by cell surface receptors. PLoS Pathogens 10(12), e1004569.

Lu, S.-W., Chen, S., Wang, J., Yu, H., Chronis, D., Mitchum, M.G. and Wang, X. (2009). Structural and functional diversity of CLAVATA3/ESR (CLE)-like genes from the potato cyst nematode Globodera rostochiensis. Molecular Plant Microbe Interactions 22, 1128–1142.

Maggenti A. R. (1981). General nematology. Springer-Verlag, New York. 372 pages.

Maier, T.R., Hewezi, T., Peng, J. and Baum, T.J. (2013). Isolation of whole esophageal gland cells from plant-parasitic nematodes for transcriptome analyses and effector identification. Molecular Plant Microbe Interactions 26, 31–35.

Mantelin, S., Thorpe, P. and Jones, J.T. (2017). Translational biology of nematode effectors. Or, to put it another way, functional analysis of effectors-what’s the point? Nematology 19, 251–261.

Mei, Y., Thorpe, P., Guzha, A., Haegeman, A., Blok, V.C., MacKenzie, K., Gheysen, G., Jones, J.T. and Mantelin, S. (2015). Only a small subset of the SPRY domain gene

Chapter 1: Introduction

family in is likely to encode effectors, two of which suppress host defences induced by the potato resistance gene. Nematology 17(4), 409–424.

Mitchum, M.G., Wang, X., Wang, J. and Davis, E.L. (2012). Role of nematode peptides and other small molecules in plant parasitism. Annual Review of Phytopathology 50, 175– 195.

Mitchum, M.G., Hussey, R.S., Baum, T.J., Wang, X., Elling, A.A., Wubben, M. and Davis, E.L. (2013). Tansley review Nematode effector proteins : an emerging paradigm of parasitism. New Phytologist 199, 879–894.

Nicol J.M., Turner S.J., Coyne D.L., Nijs L.D., Hockland S., Maafi, Z.T. (2011). Current Nematode Threats to World Agriculture. In: Genomics and Molecular Genetics of Plant- Nematode Interactions (Jones J., Gheysen G., Fenoll C., eds.), pp. 21-43. Springer, Dordrecht.

Niebel, D.A.E., Adriana, H., Paulo, F., Dirk, I., Marc, V.M. and Godelieve, G. (1996). Induction of cdc2a and cyc1At expression in Arabidopsis thaliana during early phases of nematode‐induced feeding cell formation. The Plant Journal 10, 1037–1043.

Opperman, C.H., Bird, D.M., Williamson, V.M., Rokhsar, D.S., Burke, M., Cohn, J., Cromer, J., Diener, S., Gajan, J., Graham, S., Houfek, T.D., Liu, Q., Mitros, T., Schaff, J., Schaffer, R., Scholl, E., Sosinski, B.R., Thomas, V.P. and Windham, E. (2008). Sequence and genetic map of Meloidogyne hapla: A compact nematode genome for plant parasitism. Proceedings of the National Academy of Sciences, USA 105(39), 14802–14807.

Patel, N., Hamamouch, N., Li, C., Hewezi, T., Hussey, R.S., Baum, T.J., Mitchum, M.G. and Davis, E.L. (2010). A nematode effector protein similar to annexins in host plants. Journal of Experimental Botany 61, 235–248.

Phillips, W.S., Howe, D.K., Brown, A.M.V., Eves-van den Akker, S., Dettwyler, L., Peetz, A.B., Denver, D.R. and Zasada, I.A. (2017). The draft genome of Globodera ellingtonae. Journal of Nematology 49, 127–128.

Postma, W.J., Slootweg, E.J., Rehman, S., Finkers-Tomczak, A., Tytgat, T.O.G., Van Gelderen, K., Lozano-Torres, J.L., Roosien, J., Pomp, R., Van Schaik, C., Bakker, J., Goverse, A. and Smant, G. (2012). The Effector SPRYSEC-19 of Globodera

Chapter 1: Introduction

rostochiensis suppresses CC-NB-LRR-mediated disease resistance in plants. Plant Physiology 160, 944–954.

Prior, A., Jones, J.T., Blok, V.C., Beauchamp, J., McDermott, L., Cooper, A. and Kennedy, M.W. (2001). A surface-associated retinol- and fatty acid-binding protein (Gp-FAR-1) from the potato cyst nematode Globodera pallida: lipid binding activities, structural analysis and expression pattern. Biochemical Journal 356, 387–394.

Qin, L., Overmars, H., Helder, J., Popeijus, H., Van der Voort, J.R., Groenink, W., Van Koert, P., Schots, A., Bakker, J. and Smant, G. (2000). An efficient cDNA-AFLP-based strategy for the identification of putative pathogenicity factors from the potato cyst nematode Globodera rostochiensis. Molecular Plant Microbe Interactions 13, 830–836.

Quist, C.W., Smant, G. and Helder, J. (2015). Evolution of plant parasitism in the phylum Nematoda. Annual Review of Phytopathology 53, 289–310.

Rehman, S., Postma, W., Tytgat, T., Prins, P., Qin, L., Overmars, H., Vossen, J., Spiridon, L- N., Petrescu, A-J., Goverse, A., Bakker, J. and Smant G. (2009). A secreted SPRY domain-containing protein (SPRYSEC) from the plant-parasitic nematode Globodera rostochiensis interacts with a CC-NB-LRR protein from a susceptible tomato. Molecular Plant Microbe Interactions 22, 330–340.

Rehman, S., Gupta, V.K. and Goyal, A.K. (2016). Identification and functional analysis of secreted effectors from phytoparasitic nematodes. BMC Microbiology, 16, 48.

Ritz, K. and Trudgill, D. (1999). Utility of nematode community analysis as an integrated measure of the functional state of soils: perspectives and challenges. Plant and Soil 212, 1–11.

Robertson, L., Robertson, W.M., Sobczak, M., Bakker, J., Tetaud, E., Arinagayayam, M.R., Ferguson, M.A.J., Fairlamb, A.H. and Jones, J.T. (2000). Cloning, expression and functional characterisation of a thioredoxin peroxidase from the potato cyst nematode Globodera rostochiensis. Molecular and Biochemical Parasitology 111, 41-49.

Robinson, M.P., Atkinson, H.J. and Perry, R.N. (1987). The influence of soil moisture and storage time on the motility, infectivity and lipid utilization of second stage juveniles of the potato cyst nematodes Globodera rostochiensis and G. pallida. Revue de Nematologie 10, 343–348.

Chapter 1: Introduction

Roeber, F., Jex, A.R. and Gasser, R.B. (2013). Impact of gastrointestinal parasitic nematodes of sheep, and the role of advanced molecular tools for exploring epidemiology and drug resistance - An Australian perspective. Parasites and Vectors 6, 1–13.

Rosso, M.N., Jones, J.T. and Abad, P. (2009). RNAi and functional genomics in plant parasitic nematodes. Annual Review of Phytopathology 47, 207–232.

Rosso, M.N., Hussey, R.S., Davis, E.L., Smant, G., Baum, T.J., Abad, P. and Mitchum, M.G. (2011). Nematode effector proteins: targets and functions in plant parasitism. In: Effectors in Plant–Microbe Interactions (Martin, F. and Kamoun, S., eds.), pp. 327-354. Wiley publisher.

Sasser, J.N. and Freckman, D.W. (1987). A world perspective on nematology: the role of the society. pp 7-14 in J.A. Veech and D.W. Dickson (eds.) Vistas on Nematology. Society of Nematologists, Hyattsville, Maryland.

Saucet, S.B., Cyril, V.G., Pierre, A., Henri, D. and Daniel, E. (2016). Resistance to root‐knot nematodes Meloidogyne spp. in woody plants. New Phytologist 211, 41–56.

Siddique, S., Radakovic, Z.S., De La Torre, C.M., Chronis, D., Novák, O., Ramireddy, E., Holbein, J., Matera, C., Hütten, M., Gutbrod, P., Anjam, M.S., Rozanska, E., Habash, S., Elashry, A., Sobczak, M., Kakimoto, T., Strnad, M., Schmülling, T., Mitchum, M.G. and Grundler F.M.W. (2015). A parasitic nematode releases cytokinin that controls cell division and orchestrates feeding site formation in host plants. Proceedings of the National Academy of Sciences, USA 112, 12669–12674.

Smant G., Stokkermans, J.P., Yan, Y., de Boer, J.M., Baum, T.J., Wang, X., Hussey, R.S., Gommers, F.J., Henrissat, B., Davis, E.L., Helder, J., Schots, A. and Bakker, J. (1998). Endogenous cellulases in animals: Isolation of β-1,4-endoglucanase genes from two species of plant-parasitic cyst nematodes. Proceedings of the National Academy of Sciences, USA 95(9), 4906–4911.

Smant, G. and Jones, J.T. (2011). Suppression of plant defences by nematodes. In: Genomics and Molecular Genetics of Plant-Nematode Interactions. (Jones, J., Gheysen, G., and Fenoll, C., eds.), pp. 273–286. Dordrecht: Springer Netherlands.

Sobczak, M., Golinowski, W. and Grundler, F.M.W. (1999). Ultrastructure of fedding plugs and feeding tubes formed by Heterodera schachtii. Nematology 1, 363–374.

Chapter 1: Introduction

Sobczak, M and Golinowski, W. (2011). Cyst nematode and syncytia. In: Genomics and Molecular Genetics of Plant-Nematode Interactions. (Jones, J., Gheysen, G., and Fenoll, C., eds.), pp. 61-82. Dordrecht: Springer Netherlands.

Stepek, G., Buttle, D.J., Duce, I.R. and Behnke, J.M. (2006). Human gastrointestinal nematode infections: Are new control methods required? International Journal of Experimental Pathology 87, 325–341.

Szitenberg, A., Salazar-Jaramillo, L., Blok, V.C., Laetsch, D.R., Joseph, S., Williamson, V.M., Blaxter, M.L. and Lunt, D.H. (2017). Comparative genomics of apomictic root- knot nematodes: Hybridization, ploidy, and dynamic genome change. Genome Biology and Evolution 9(10), 2844–2861.

The C. elegans Sequencing Consortium (1998). Genome sequence of the nematode C. elegans : A platform for investigating biology. Science 282(5396), 2012–2018.

Thorpe, P., Mantelin, S., Cock, P.J.A., Blok, V.C., Coke, M.C., Eves van den Akker, S., Guzeeva, E.A., Lilley, C.J., Smant, G., Reid, A.J., Wright, K.M., Urwin, P.E. and Jones J.T. (2014). Genomic characterisation of the effector complement of the potato cyst nematode Globodera pallida. BMC Genomics 15, 923.

Turner, S.J. and Rowe, J.A. (2006). Cyst nematodes. In: Plant nematology (Perry, R.N. and Moens, M., eds.), pp. 90–122. Wallingford, Oxfordshire: CAB International.

Ugarte, C. and Zaborski, E. (2014). Soil nematodes in organic farming systems. 1–11.

Urwin, P.E., Lilley, C.J. and Atkinson, H.J. (2002). Ingestion of double-stranded RNA by preparasitic juvenile cyst nematodes leads to RNA interference. Molecular Plant- Microbe Interaction 15, 747–752.

Vanholme, B., De Meutter, J., Tytgat, T., Van-Montagu, M., Coomans, A. and Gheysen, G. (2004). Secretions of plant-parasitic nematodes: A molecular update. Gene 332, 13–27.

Van Megen, H., Van Den Elsen, S., Holterman, M., Karssen, G., Mooyman, P., Bongers, T., Holovachov, O., Bakker, J. and Helder, J. (2009). A phylogenetic tree of nematodes based on about 1200 full-length small subunit ribosomal DNA sequences. Nematology 11, 927–950.

Vieira, P., Eves-Van Den Akker, S., Verma, R., Wantoch, S., Eisenback, J.D. and Kamo, K. (2015). The pratylenchus penetrans transcriptome as a source for the development of

Chapter 1: Introduction

alternative control strategies: Mining for putative genes involved in parasitism and evaluation of in planta RNAi. PLoS One 10, 1–25.

Wang, X., Mitchum, M.G., Gao, B., Li, C., Diab, H., Baum, T.J., Hussey, R.S. and Davis, E.L. (2005). A parasitism gene from a plant-parasitic nematode with function similar to CLAVATA3/ESR (CLE) of Arabidopsis thaliana. Molecular Plant Pathology 6, 187– 191.

Wang, J., Lee, C., Replogle, A., Joshi, S., Korkin, D., Hussey, R., Baum, T.J., Davis, E.L., Wang, X. and Mitchum, M.G. (2010). Dual roles for the variable domain in protein trafficking and host-specific recognition of Heterodera glycines CLE effector proteins. New Phytologist 187, 1003–1017.

WHO (2013). WHO-Onchocerciasis. (http://www.who.int/onchocerciasis/en/).

Wieczorek, K., Golecki, B., Gerdes, L., Heinen, P., Szakasits, D., Durachko, D.M., Cosgrove, D.J., Kreil, D.P., Puzio, P.S., Bohlmann, H. and Grundler, F.M.W. (2006). Expansins are involved in the formation of nematode-induced syncytia in roots of Arabidopsis thaliana. The Plant Journal 48, 98–112.

Williamson, V.M. and Hussey, R.S. (1996). Nematode pathogenesis and resistance in plants. Plant Cell 8, 1735.

Williamson, V.M. and Gleason, C.A. (2003). Plant-nematode interactions. Current Opinion in Plant Biology 6, 327–333.

Wyss, U. and Zunke, U. (1986). Observation of the behaviour of second stage juveniles of Heterodera schachtii inside host roots. Revue de Nématologie 9, 153–166.

Wyss, U. (1992). Observations on the feeding behavior of Heterodera schachtii throughout development, including events during moulting. Fundamentals of Applied Nematology 15, 75–89.

Xue, B., Hamamouch, N., Li, C., Huang, G., Hussey, R.S., Baum, T.J. and Davis, E.L. (2013). The 8D05 parasitism gene of Meloidogyne incognita is required for successful infection of host roots. Phytopathology 103, 175–81.

Yadav, B.C., Veluthambi, K. and Subramaniam, K. (2006). Host-generated double stranded RNA induces RNAi in plant-parasitic nematodes and protects the host from infection. Molecular and Biochemical Parasitology 148, 219–222.

Chapter 1: Introduction

Yang, D., Chen, C., Liu, Q. and Jian, H. (2017). Comparative analysis of pre- and post- parasitic transcriptomes and mining pioneer effectors of Heterodera avenae. Cell Bioscience 7, 11.

Zhang, Z.Q. (2013). Animal biodiversity: An update of classification and diversity. Zootaxa 3703, 5–11.

Zheng, J., Peng, D., Chen, L., Liu, H., Chen, F., Xu, M., Ju, S., Ruan, L. and Sun, M. (2016). The Ditylenchus destructor genome provides new insights into the evolution of plant parasitic nematodes. Proceedings of the Royal Society B: Biological Sciences 283,1835.

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

Somnath S. Pokhare1,2, Samer S. Habash1, John T. Jones3, Abdelnaser Elashry1,4 & Florian M.W. Grundler1

1 INRES Molecular Phytomedicine, Rheinische-Friedrich-Wilhelms-University of Bonn, D- 53115 Bonn, Germany.

2 Crop Protection, ICAR-National Rice Research Institute, Cuttack-753006, Odisha, India.

3 Cell and Molecular Sciences Group, Dundee Effector Consortium, James Hutton Institute, Dundee DD2 5DA, UK.

4 Current address: Strube Research GmbH & Co. KG, Hauptstraße 1, 38387, Söllingen, Germany.

*Manuscripts accepted to Nematology journal for publication

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

2.1 Summary

This paper presents studies on the life cycle of Heterodera sacchari under in vitro conditions. Pluronic gel was used as a medium for growth of H. sacchari. The life cycle was completed in 7-9 weeks on rice (Oryza sativa, ‘Nipponbare’). After infection, juveniles developed and reached the reproducing adult female stage at 25 days post inoculation (dpi). At 35 dpi, all females produced eggs in various numbers. Some females were translucent, and eggs inside could be counted. At 49 dpi females started to tan and developed into dark brown cysts.

Hatching of H. sacchari juveniles from cysts could be stimulated by 3mM ZnCl2 but not by rice root exudates. The in vitro culture of H. sacchari on pluronic gel can be used efficiently to collect post-infective nematode/host samples at different time points for various studies and to screen different rice cultivars for resistance/susceptibility.

Keywords - cyst nematodes, ‘Nipponbare’, Oryza sativa, plant–parasitic nematodes, rice, sugarcane cyst nematode.

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

2.2 Introduction

Rice is the world’s most important staple food crop and is the primary source of carbohydrate for more than half of the world’s population (Somado et al., 2008). Two domesticated species of rice are grown: Oryza sativa, commonly known as Asian rice, is cultivated throughout the world, whilst O. glaberrima (African rice) is grown extensively in Africa. Productivity of rice varies from less than 1 t ha-1 under poorly managed, rainfed conditions to more than 10 t ha-1 under well-managed irrigated systems following good agricultural practices (GRiSP, 2013). Global demand for rice is increasing and is expected to reach 555 million tons by 2035 due to the increasing human population (Seck et al., 2012). Increasing rice yields is of key importance for achieving global food security. Reducing or avoiding losses that occur during and after production due to pests and diseases has a significant role to play in this process. On average, farmers lose 37% of their produce due to pests and diseases including insects, microbial diseases, weeds and nematodes (Anonymous, 2018).

Plant–parasitic nematodes can cause annual losses of up to 25% in rice production (Bridge et al., 2005; Kyndt et al., 2014). Over 150 species of plant–parasitic nematodes infect rice including root parasites, such as root-knot nematodes of the genus Meloidogyne, cyst nematodes belonging to Heterodera, root lesion nematode Pratylenchus spp. and rice root nematode, Hirschmanniella spp. Shoot parasites (foliar parasites) include rice stem nematode, Ditylenchus angustus, and rice leaf nematode, Aphelenchoides besseyi, causing ufra and white tip disease, respectively (Fortuner & Merny 1979; Catling & Islam 1999).

Although extensive studies of the interactions between root-knot nematodes (particularly M. graminicola) and rice have been carried out (Haegeman et al., 2013; Petitot et al., 2017), little information is currently available on interactions between rice and cyst nematodes. Four different species of cyst nematodes infect rice roots under both upland and irrigated conditions. These are H. oryzae, H. oryzicola, H. elachista and H. sacchari (Lorieux et al., 2003; Bridge et al., 2005). Heterodera oryzicola and H. elachista have a relatively restricted distribution, whilst H. oryzae and H. sacchari are more widespread (Jayaprakash & Rao 1982; De Luca et al., 2013).

Heterodera sacchari is commonly known as the sugarcane cyst nematode as it was reported for the first time as a parasite of sugarcane (Saccharum officinale) in Congo-Brazzaville (Luc & Merny, 1963). This nematode was subsequently also described as a parasite of rice (Merny, 1970). To date, only rice and sugarcane are reported as primary cultivated hosts of

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

this nematode, but it also infects and multiplies on several weed hosts of the poaceae / gramineae family. Heterodera sacchari has been reported from Africa (Benin, Burkina Faso, Chad, Congo, Côte d’Ivoire, Ghana, Liberia, Nigeria, Senegal and The Gambia), from Asia (India, Pakistan and Thailand) and from Jamaica (Anonymous, 2014). Although H. sacchari is reported in both upland and lowland rice cultivations, it has a greater impact under upland conditions (Coyne & Plowright, 2000; Bridge, 2005).

Symptoms of H. sacchari are similar to those of other cyst nematodes in that they are nonspecific and similar to those of plants that are nutrient deficient. These symptoms include severe leaf chlorosis, short and necrotic roots, poor plant vigour with lower tiller numbers, delayed panicle emergence and poor grain filling, resulting in significant yield reduction (Babatola, 1983; Akpheokhai & Claudius-Cole, 2017). Screening for resistance against H. sacchari has shown that all forty-three O. sativa cultivars tested were highly susceptible, while fifteen out of twenty-one accessions of O. glaberrima and seven of the nine accessions of O. breviligulata were resistant (Reversat & Destombes, 1998). In another study, Plowright et al. (1999) found that all O. glaberrima lines were resistant to two isolates (Côte d’Ivoire and Ghana) of H. sacchari. One O. glaberrima accession, ‘TOG5681’ was used as a resistance donor for H. sacchari and resistance was found to be controlled by one major gene, Hsa-1Og, with codominance of the susceptible and resistant alleles (Lorieux et al., 2003).

Detailed understanding of the interaction between plant-parasitic nematodes and their hosts is critical for the development of new control strategies. For any pathosystem an effective culture system will facilitate the study of nematode behaviour, including the ability of pre- parasitic juveniles to locate host roots, development of the nematode and response of the host to infection. Although life cycle studies using monoxenic cultures can be accomplished in pots using soil or sand adsorbent polymer (SAP) substrates, in vitro methods with agar plates or in pluronic gel procedures offer the potential for continuous and detailed observations.

In vitro assays are well established to study infection and development for various cyst nematodes (H. schachtii, H. trifolii and H. cajani), root-knot nematodes (M. incognita and M. arenaria) and lesion nematode (P. penetrans) on Arabidopsis thaliana in Knop medium (Sijmons et al., 1991). Currently, the sugar beet cyst nematode, H. schachtii, infecting A. thaliana serves as a model to study the host nematode interaction at the molecular level (Sijmons et al., 1991). There have also been a number of studies using pluronic F-127 as medium for nematode-plant interactions (Wang et al., 2009a, b; Dutta et al., 2011; Sasaki- Crawley et al., 2012). Pluronic F-127 gel is a non-ionic surfactant, co-polymer of propylene

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

oxide and ethylene oxide (Rokhade et al., 2007). Pluronic gel has been used as medium to study M. incognita behaviour and attraction towards tomato, common bean and Arabidopsis (Wang et al., 2009a), and also to compare host recognition, invasion, development and reproduction of M. incognita and M. graminicola on rice and tomato (Dutta et al., 2011). In another study, rice cultivars were screened for resistance to M. graminicola and to compare infection and development of nematode on resistant and susceptible rice cultivars (Kumari et al., 2016).

Here we describe a protocol for culture and observation of H. sacchari on rice using pluronic gel as a medium and use this system to analyse the life cycle of H. sacchari in detail.

2.3 Materials and methods

Nematode culture

Four different rice cultivars, ‘IR64’, ‘Nipponbare’, ‘MTU9’ and ‘TN1’ (all O. sativa type) were tested to establish their levels of susceptibility to H. sacchari under glasshouse conditions. Seeds of these cultivars were soaked overnight in sterile distilled water and kept on sterile wet filter paper inside Petri plates at 250C and 16 h: 8 h light: dark photoperiod to allow germination. After 1-week, germinated seeds were transplanted to pots containing sterilised potting mixture of sand, field soil and organic matter (70:29:1). For each cultivar ten replicates were used to test susceptibility. Cysts of H. sacchari were surface sterilised with 0.5% sodium hypochlorite (NaOCl) for 10 min and incubated in water followed by

3mM zinc chloride (ZnCl2) solution to initiate hatching. Freshly hatched pre-parasitic juveniles (J2) were used for infection. Potted plants were inoculated with 200 J2 2 weeks after transplanting and watered every 2 days and fertilised with water soluble Nitrophoska once in 15 days. Twelve weeks after inoculation, watering was stopped for 2 weeks to allow the plants to dry. Soil was collected and washed with 250 mm sieves and supernatant was collected in a beaker to check presence of cysts. Number of cysts (250 g soil)-1 were counted and significant differences were determined based on Fishers LSD method (P < 0.05). As ‘Nipponbare’ is a widely accepted model rice cultivar, we used this cultivar for all further studies.

Hatching assay

Heterodera sacchari cysts were surface sterilised after harvesting as described above. One- week-old rice seedlings were used to prepare root exudates. For this, four plants were transferred to a small flask containing 50 ml of distilled water and the roots were submerged

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

in the water. The water was collected after 24 h, filtered with 0.2 µm filter and used as rice root exudate. The hatching assay was done using distilled water, 3mM ZnCl2 and rice root exudates in 24 well tissue culture plates. One ml of each solution was added per well followed by 25 surface sterilised cyst. Each treatment had five replicates with 25 cysts per replicate. The total numbers of hatched J2 were counted at 4-day intervals until no significant changes in numbers were observed in two consecutive observations. Data were analysed using one-way ANOVA and all pairwise multiple comparison procedures used Fisher’s LSD method) (P < 0.05).

Preparation, sterilisation and germination of rice seeds

‘Nipponbare’ seeds were surface sterilized by soaking overnight in sterile distilled water on a shaker followed by treatment with 70% ethanol and 6% NaOCl each for 15 min. The previously described method using 70% ethanol for 30 s after overnight soaking in distilled water (Kumari et al., 2016) failed to sterilise seeds effectively under the conditions tested here. Treated seeds were washed four times (2 min each) with sterile water and seeded on media plates (Knop and MS medium) or spread on sterile wet filter paper in a Petri dish for germination and incubated at 25°C. One week-old seedlings were transplanted from wet filter paper onto the Petri plates with pluronic gel liquid under sterile conditions. Plates were sealed with parafilm when pluronic solution had solidified. Pluronic gel F-127 was prepared by adding 23 g of Pluronic F-127 powder (Sigma) to 80 ml of double distilled water (Wang et al., 2009a). The resulting gel was sterilised in an autoclave set at 121°C. After autoclaving, the sterilised gel was kept at 10°C overnight on a shaker to allow it to dissolve and then stored at 10°C. Aliquots of this were used for all the in vitro experiments.

Plant infection and life cycle observations

Cysts were harvested from ‘Nipponbare’ plants grown in pots in the glasshouse 12 weeks after inoculation. Cysts were collected and surface sterilised using 0.5% NaOCl for 10 min and rinsed four times with distilled water (Pariyar et al., 2016). After sterilisation, cysts were incubated in 3 mM ZnCl2 to facilitate hatching of J2. Freshly hatched J2 were collected every 5 days and surface sterilised using 0.05% mercuric chloride for 4 min. Around 150 J2 (plant)- 1 were used to inoculate plants 1 day after they had been transplanted onto the pluronic gel. Infected plates were incubated in a growth chamber with 16 h light and 8 h dark at 25°C and checked at different time points under binocular microscope to observe the development of the nematodes on the infected roots. This was documented by staining the infected roots with

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

acid fuchsin during early stages of infection (Byrd et al., 1983) or taking images of the nematodes either inside the root tissues or attached to the roots using Leica M165C Binocular (Leica Microsystems) and Leica Application Suite software.

2.4 Results

Optimization of culture conditions

All the tested rice cultivars were found to be susceptible to H. sacchari with the highest number of cysts found in pots planted with ‘Nipponbare’ plants (Fig. 2.1).

Hatching assay

During the hatching assay, we found that 3 mM ZnCl2 initiated hatching and that the total number of hatched J2 in ZnCl2 was significantly higher than those emerging in water or rice root exudates at 8, 12 and 16 days after exposure to the hatching agents. The hatching of J2 from the cysts exposed to rice root exudates was not significantly higher at any of the time points tested. However, by 20 days after exposure to hatching agents, no significant differences between any of the treatments were observed (Fig. 2.2). Exposure to ZnCl2 may therefore increase the initial rate of hatch of H. sacchari.

Development of H. sacchari on rice under in vitro conditions

During initial experiments, we found that H. sacchari J2 did not infect rice grown on Knop and MS media efficiently. After inoculating 150 J2 plant-1, we found a maximum of 2-3 developed females plant-1 in just 4 out of 10 plants, with another 6 plants showing no development of nematodes. By contrast, nematodes developed well on plants grown in the pluronic gel.

Nematodes were attracted to rice plant roots grown in the pluronic gel. Within 24 h, J2 were able to locate the roots and started browsing near the root tips. A slight root swelling was observed at the point of nematode infection (Fig. 2.3A). Nematodes were observed with the anterior part of their body inside the root tissues after staining infected roots with acid fuchsin at 3 days post infection (dpi) (Fig. 2.3B) and 7 dpi (Fig. 2.3C). After 10-12 dpi, nematodes that had completely entered the roots started to protrude from the root tissues (Fig. 2.3D). Young white females were observed attached to the root at 15 dpi with syncytia clearly visible (Fig. 2.3E). In the vast majority of cases the developed nematodes became females. Males were very rarely observed at 20-25 dpi but only when very heavy inoculations (>300 J2 plant-1) were used. The presence of males was not required for further development of

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

eggs, in line with previous observations that this nematode reproduces by mitotic parthenogenesis (Lorieux et al., 2003; Anonymous, 2014). Egg production started at around 25 dpi, and eggs were visible through the translucent female nematodes (Fig. 2.3F). The numbers of eggs varied enormously in between females (Fig. 2.3G, H). At 40 dpi, females stated to tan and they became darker as the life cycle progressed. Around 50 days after inoculation dark brown cysts were observed (Fig. 2.3I). From the 9th week of inoculation onwards cysts were picked and crushed in a drop of water to examine the eggs inside. Fully developed eggs inside the cysts were detected (not shown).

2.5 Discussion

The main aim of this work was to develop a system for in vitro culture of H. sacchari that allowed visualisation of the various developmental stages of the nematode. This allows collection of various life stages for transcriptome analysis as well as screening for resistance and other life cycle studies.

Some cyst nematodes coordinate their life cycles with those of their hosts by hatching solely in response to root exudates of host plants although others, most frequently those with a broader host range, hatch freely in water (Masler & Perry, 2018). ZnCl2 is known to stimulate hatch in many species that do not require exposure to host diffusates (Sijmons et al., 1991). We observed that the hatching of H. sacchari did not respond to rice root exudates to any greater extent than water, but it showed an acceleration of hatching in the presence of ZnCl2. This maybe a reflection of the relatively broad host range of H. sacchari but it is also possible that the nematode may respond differently to diffusates of other host species.

We found that pluronic gel was a suitable medium for growing rice plants that could support H. sacchari to develop through the complete life cycle. The main advantage of using pluronic gel is its transparency, which makes it possible continuously to observe nematode movement as well as development inside the root (Wang et al., 2009a; Kumari et al., 2016). In addition, the gel is in a liquid state at temperatures below 10oC, allowing nematodes to be applied as a suspension, and thus synchronising the infection process, as well as allowing easy collection of clean parasitic life stages or infected plant materials for further analysis under sterile conditions without contamination with soil (Gardener & Jones 1984). Heterodera sacchari completed its life cycle within 7-9 weeks on pluronic gel with all nematode developmental stages observed on plants. As previously described for M. graminicola (Kumari et al., 2016),

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

the pluronic gel system is therefore well suited for assays that aim to determine susceptibility or resistance of rice cultivars.

Acknowledgements

Somnath S. Pokhare is supported by a fellowship provided by the Indian Council of Agricultural Research (ICAR), Government of India. The James Hutton Institute receives funding from the Scottish Government. This work is also benefited from interactions funded by Royal Society International Exchanges project IE150670. Authors also thank to Prof. Michael Frei (INRES, Plant nutrition) for providing rice seeds and Danny Coyne (IITA, Ibadan, Nigeria) for nematode population.

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

2.6 Figures

1600

1 *

- 1400 1200 1000 800 600 400

No. of cysts (250 cysts soil)g of No. 200 0 'IR64' 'TN1' 'MTU9' 'Nipponbare' Rice cultivar

Fig. 2.1: Levels of susceptibility of different rice varieties to Heterodera sacchari under glasshouse conditions. Ten plants of each variety were transplanted to pots in the glasshouse. 200 second-stage juveniles plant-1 were inoculated 2 weeks after transplanting. Number of cysts (250 g soil)-1 were counted 12 weeks after inoculation. Asterisks indicate significant differences based on Fishers LSD method (P < 0.05).

1400 1200 1000 * 800 * 600 * 400

Total no. of hatched J2hatched of no. Total 200 0 4 days 8 days 12 days 16 days 20 days Time point of observation

Water Zinc Chloride Rice Exudates

Fig. 2.2: Heterodera sacchari hatching assay using water, 3 mM ZnCl2 and rice root exudates. Number of cysts per treatment = 25. No. of replicates = 5. Interval of counting hatched second-stage juveniles (J2) = 4 days. Data are normally distributed and had equal

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

variance. Data were analysed using one-way ANOVA and all pairwise multiple comparison procedures used Fisher’s LSD method (P < 0.05).

Fig. 2.3: In vitro life cycle of Heterodera sacchari on pluronic gel. A: Slight root swelling at the infection point of second-stage juveniles. B, C: Acid fuchsin staining of infected nematodes at 3 and 7 days post inoculation (dpi). D: Developing nematodes with root ruptured at 10-12 dpi. E: Young H. sacchari female at 15 dpi with visible syncytium. F: Translucent female with eggs inside at 25 dpi. G, H: Variation in egg number in developing females at 35-40 dpi. I: Dark brown cysts after 50 dpi.

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

2.7 Reference

Akpheokhai, L.I. & Claudius-Cole, A.O. (2017). Effect of Heterodera sacchari on growth and yield of selected upland NERICA rice in Nigeria. Tropical plant pathology 42(6), 431-442.

Annonymous (2014) CABI’s report on sugarcane cyst nematode, Heterodera sacchari. (http://download.ceris.purdue.edu/file/3047).

Annonymus (2018). IRRI Rice Knowledge Bank (http://www.knowledgebank.irri.org/step- by-step-production/growth/pests-and-diseases).

Babatola, J.O. (1983). Pathogenicity of Heterodera sacchari on rice. Nematologia Mediterranea 11, 21–25.

Bridge, J., Plowright, R.A. and DeLiang, Peng. (2005). Nematode parasites of rice. In: Luc, M., Sikora, R.A. & Bridge, J. (eds.). Plant parasitic nematodes in subtropical and tropical agriculture. CABI, pp. 87–128.

Byrd, D.W., Kirkpatrick, T. and Barker, K.R. (1983). An improved technique for clearing and staining plant tissue for detection of nematodes. Journal of Nematology 15, 142- 143.

Catling, H.D. and Islam, Z. (1999). Pests of deepwater rice and their management. Integrated Pest Management Reviews 4, 193–229.

Coyne, D.L. and Plowright, R.A. (2000). Pathogenicity of cyst nematode, Heterodera sacchari, on rice in sand and clay soil. International Rice Research Notes 25 (1), 17–18.

De Luca, F., Vovlas, N., Lucarelli, G., Troccoli, A., Radicci, V., Fanelli, E., Cantalapiedra- Navarrete, C., Palomares-Rius, J.E. and Castillo, P. (2013). Heterodera elachista the Japanese cyst nematode parasitizing corn in northern Italy: Integrative diagnosis and bionomics. European Journal of Plant Pathology 136, 857–872.

Dutta, T.K., Powers, S.J., Kerry, B.R., Gaur, H.S. and Curtis, R.H.C. (2011). Comparison of host recognition, invasion, development and reproduction of Meloidogyne graminicola and M. incognita on rice and tomato. Nematology 13, 509–520.

Fortuner, R. and Merny, G. (1979). Root-parasitic nematodes of rice. Nematology 2, 79–102.

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

Gardener, S. and Jones, J.G. (1984). A new solidifying agent for culture media which liquefies on cooling. Journal of General Microbiology 130, 731–733.

GRiSP (2013). Global Rice Science Partnership. GRiSP in motion, Annual Report 2012. Los Baños (Philippines), International Rice Research Institute, pp. 32.

Jayaprakash, A. and Rao, Y.S. (1982). Life history and behaviour of the cyst nematode, Heterodera oryzicola Rao and Jayaprakash, 1978 in rice (Oryza sativa L.). Proceedings of the Indian Academy of Sciences. Section B- Biological Sciences 91, 283-295.

Kumari, C., Dutta, T.K., Banakar, P. and Rao, U. (2016). Comparing the defence-related gene expression changes upon root-knot nematode attack in susceptible versus resistant cultivars of rice. Scientific Reports 6, 22846.

Kyndt, T., Fernandez, D. and Gheysen, G. (2014). Plant-parasitic nematode infections in rice: Molecular and cellular insights. Annual Review of Phytopathology 52, 135–153.

Lorieux, M., Reversat, G., Garcia Diaz, S.X., Denance, C., Jouvenet, N., Orieux, Y., Bourger, N., Pando-Bahuon, A. and Ghesquière, A. (2003). Linkage mapping of Hsa-10g, a resistance gene of African rice to the cyst nematode, Heterodera sacchari. Theoretical and Applied Genetics 107, 691–696.

Luc, M. and Merny, G. (1963). Heterodera sacchari N. SP. (Nematoda: Tylenchoidea) parasite de la canne a sucre au Congo-Brazzaville. Nematologica 9, 31-37.

Masler, E.P. and Perry, R.N. (2018) Hatch, survival and sensory perception. In: Perry, R.N., Moens, M., Jones, J.T. (eds.). Cyst nematodes. CABI, pp. 44-73.

Merny, G. (1970). Les nématodes phytoparasites des rizières inondées en Côte d'Ivoire. 1. Les espèces observées. Cahiers ORSTOM.Série Biologie, (11 spécial "Nématologie"), pp.3-43.

Pariyar, S.R., Dababat, A.A., Siddique, S., Erginbas-Orakci, G., Elashry, A., Morgounov, A. and Grundler, F.M.W. (2016). Identification and characterisation of resistance to the cereal cyst nematode Heterodera filipjevi in winter wheat. Nematology 18, 377–402.

Chapter 2: In vitro life cycle of Heterodera sacchari on pluronic gel

Petitot, A.S., Kyndt, T., Haidar, R., Dereeper, A., Collin, M., De Almeida Engler, J., Gheysen, G. and Fernandez, D. (2017). Transcriptomic and histological responses of African rice (Oryza glaberrima) to Meloidogyne graminicola provide new insights into root-knot nematode resistance in monocots. Annals of Botany 119(5), 885–899.

Plowright, R.A., Coyne, D.L., Nash, P. and Jones, M.P. (1999). Resistance to the rice nematodes Heterodera sacchari, Meloidogyne graminicola and M . incognita in Oryza glaberrima and O. glaberrima X O. sativa interspefic hybrids. Nematology 1, 745–751.

Reversat, G. and Destombes, D. (1998). Screening for resistance to Heterodera sacchari in the two cultivated rice species, Oryza sativa and O. glaberrima. Fundamental and Applied Nematology 21, 307–317.

Rokhade, A., Shelke, N., Patil, S. and Aminabhavi, T. (2007). Novel hydrogel microspheres of chitosan and pluronic F-127 for controlled release of 5-fluorouracil. Journal of Microencapsulation 24, 274–288.

Sasaki-Crawley, A., Curtis, R., Birkett, M., Papadopoulos, A., Blackshaw, R. and Pickett, J. (2012). The use of Pluronic F-127 to study the development of the potato cyst nematode, Globodera pallida. Nematology 14, 869–873.

Seck, P.A., Diagne, A., Mohanty, S. and Wopereis, M.C.S. (2012). Crops that feed the world 7: Rice. Food Security 4, 7–24.

Sijmons, P.C., Grundler, F.M.W., Von Mende, N., Burrows, P.R. and Wyss, U. (1991). Arabidopsis thaliana as a new model host for plant‐parasitic nematodes. The Plant Journal 1, 245–254.

Somado, E.A., Guei, R.G. and Nguyen, N. (2008). Overview: Rice in Africa In: Somado, E.A., Guei, R.G. & Keya, S.O. (eds.). NERICA: the New Rice for Africa – a Compendium. Africa Rice Center (WARDA), pp. 1-9.

Wang, C., Lower, S. and Williamson, V.M. (2009a). Application of pluronic gel to the study of root-knot nematode behaviour. Nematology 11, 453–464.

Wang, C., Bruening, G. and Williamson, V.M. (2009b). Determination of preferred pH for root-knot nematode aggregation using pluronic F-127 gel. Journal of Chemical Ecology 35, 1242–1251.

Chapter 3: Signatures of adaptation to a monocot host in the plant-parasitic nematode Heterodera sacchari

Somnath S. Pokhare1,2, Peter Thorpe3, Sebastian Eves-van den Akker4, Pete Hedley3, Samer S. Habash1, Abdelnaser Elashry1*, Florian M.W. Grundler1 & John T. Jones3,5

1Institute of Crop Science and Resource Conservation, Department of Molecular Phytomedicine, University of Bonn, 53115 Bonn, Germany. 2Crop Protection Division, ICAR-National Rice Research Institute, Cuttack 753006, Odisha, India. 3The James Hutton Institute, Invergowrie, Dundee, DD2 5DA, UK. 4Department of Plant Sciences, University of Cambridge, Cambridge, CB2 3EA. 5School of Biology, University of St Andrews, North Haugh, St Andrews, KY16 9TZ.

*Current address: Strube Research GmbH & Co. KG, Hauptstraße 1, 38387, Söllingen, Germany.

* Manuscript in preparation

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

3.1 Abstract Most major food crops of the world are threatened by at least one species of parasitic nematode. Given that food crops are a relatively modern development, the current state is at least in part the result of adaptation of the parasite. Understanding the genes involved in adaptation to a new host is of fundamental importance. Interactions between plant-parasitic nematodes and their host plants are mediated by effectors; secreted proteins and small molecules that manipulate the plant to the benefit of the pathogen. To understand the role of plant-parasitic nematode effectors in host adaptation, we sequenced and analysed the transcriptome of an increasingly economically important cyst nematode parasite of rice and sugarcane, Heterodera sacchari.

We use a multi-gene phylogenetic approach to show that H. sacchari and the related cereal cyst nematode Heterodera avenae share a common evolutionary origin, and that they evolved to parasitise monocotyledonous plants from a last common dicotyledon-parasitic ancestor. We compare the cell wall degrading enzyme and effector repertoires of the derived monocotyledonous plant-parasite with those of the dicotyledonous parasites H. glycines and Globodera rostochiensis to better understand the consequences of this transition. We show that, in general, the effector repertoires are similar between the two species. A comparison of effectors and non-effectors between H. sacchari and G. rostochiensis suggests that the effectors have accumulated more mutations than non-effectors since these species diverged. While most effectors show conserved spatiotemporal expression profiles and likely function, specific aspects of the effector repertoire of H. sacchari may be adapted to monocotyledonous plants. This is exemplified by the plant-peptide hormone mimics, the CLAVATA3/EMBRYO SURROUNDING REGION-like (CLE) effectors. We show that the peptide hormones encoded by H. sacchari CLE effectors are more similar to those from rice than those from other plants, or those from other plant-parasitic nematodes. Finally, we experimentally validate the functional significance of these observations by demonstrating that CLE peptides encoded by H. sacchari induce a short root phenotype in rice, whereas CLE peptides encoded by a related parasite of dicotyledonous plants do not. These data provide a functional example of effector evolution that co-occurred with the transition from a dicotyledonous-parasitic to a monocotyledonous-parasitic lifestyle.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

3.2 Introduction

Plant-parasitic nematodes cause damage to world agriculture valued at approximately $80 billion each year (Nicol et al., 2011; Jones et al., 2013). However, many growers (particularly in developing nations) are unaware of the damage caused by plant-parasitic nematode because they are small, soil-dwelling and cause non-specific symptoms. The true extent of the damage caused by plant-parasitic nematodes is therefore likely to be considerably higher than this. The cyst forming nematodes (Heterodera and Globodera spp.) are one of the most damaging plant-parasitic nematode groups. These sedentary endoparasitic pathogens are obligate biotrophs and induce the formation of a large multinucleate, metabolically active syncytium in the roots of their hosts (reviewed in Perry, Moens & Jones, 2018). The syncytium is formed by progressive fusion of cells from an initial syncytial cell selected by the nematode after invasion of the host (Sobczak & Golinowski, 2011). Syncytium formation is associated with profound changes in host gene expression and modulation of the cell cycle (e.g. Gheysen & Mitchum, 2011).

The interactions between plants and their pathogens, including cyst nematodes, are mediated by effectors; secreted proteins that manipulate the host to the benefit of the pathogen. Effectors from cyst nematodes are primarily produced in the dorsal and subventral pharyngeal gland cells and are secreted into the host via the stylet. The availability of genome and/or transcriptome resources from a range of cyst nematodes (e.g. Gao et al., 2003; Cotton et al., 2014; Kumar et al., 2014; Zheng et al., 2015; Eves-van den Akker et al., 2016) has facilitated identification of effectors from these species. Strategies for identifying effectors from genome and transcriptome resources include identifying secreted proteins that are upregulated at parasitic stages of the nematode (e.g. Thorpe et al., 2014; Espada et al., 2016) or direct sequencing of mRNA extracted from aspirated gland cells (Maier et al., 2013) followed in both cases by in situ hybridisation to confirm expression in the pharyngeal gland cells. More recently, it has been shown that promoters associated with genes expressed in the gland cells can be used to identify comprehensive lists of effectors from diverse PPN species (Eves-van den Akker & Birch, 2016; Espada et al., 2018). As a result of these studies, effectors have been identified from a wide range of PPN with subsequent functional studies showing that they have roles in various stages of the plant-nematode interaction including metabolism of the plant cell wall to facilitate invasion and migration, suppression of host defences and initiation of the syncytium (reviewed in Jones & Mitchum, 2018).

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

The plant cell wall is the first significant barrier that any invading pathogen, including plant- parasitic nematodes, will need to overcome to infect a plant and plant-parasitic nematodes are well equipped with proteins that allow them to metabolise the plant cell wall. The first effector identified from any plant-parasitic nematode was a beta 1,4 endoglucanase (cellulase) from the potato cyst nematode Globodera rostochiensis (Smant et al., 1998) and a range of cell wall degrading and modifying proteins have subsequently been identified as cyst nematode effectors including pectate lyase (Popeijus et al., 2000), GHF43 Arabinase (Cotton et al., 2014), GH53 Arabinogalactan endo- 1,4 beta galactosidase (Vanholme et al., 2009), expansins (Qin et al., 2004) and proteins encoding carbohydrate binding domains (Hewezi et al., 2008). All of these genes, as well as others encoding chorismate mutase (Jones et al., 2003) and proteins potentially involved in vitamin biosynthesis (Craig et al., 2008), have been acquired by horizontal gene transfer from bacteria (reviewed in Kikuchi et al., 2017). Cyst nematode effectors have also been identified that suppress host defence responses, most notably several members of the SPRYSEC family of effectors (Postma et al., 2012; Mei et al., 2015) and a modified ubiquitin extension protein (Chronis et al., 2013). The details of how cyst nematodes induce the formation of their syncytium in the roots of their hosts are less clear, although effectors that are likely to be important in this process have been characterised. A novel effector (19C07) has been identified from Heterodera glycines and H. schachtii that interacts with the LAX3 auxin influx transporter (Lee et al., 2011). In addition, all cyst nematodes studied to date produce effectors that include variable numbers of C- terminal repeats encoding peptides similar to CLAVATA3/EMBRYO SURROUNDING REGION-like (CLE) peptides. Functional studies have shown that the nematode peptides can complement mutant Arabidopsis lacking these peptides (Wang et al., 2005). The CLE proteins are modified by plant cell machinery in a manner similar to that of the endogenous proteins and the CLE peptides themselves subsequently interact with the CLAVATA2 receptor protein, which is required for nematode parasitism (Replogle et al., 2011; Replogle et al., 2013). In plants, CLE peptides regulate cell differentiation and thus contribute to the control of meristem maintenance in shoots, roots and vascular tissues, and the ability to produce endogenous CLE peptides is likely be a key factor in the ability to induce a feeding structure in plants.

A very wide range of plant species are parasitized by cyst nematodes, including monocots and dicots. However, each individual species of cyst nematode tends to have relatively narrow host range, with some exceptions. Although very little is known about the molecular

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari determinants of host range in plant parasitic nematodes, effectors have been shown to have a central role in this process in other pathogens (reviewed by Stam et al., 2014). A view has emerged “non-host resistance” of closely related species is most likely due to recognition of effectors by a resistance gene, while failure to infect a more distantly related species is most likely due to the incompatibility of effectors with their cognate targets (Schulze-Lefert & Panstruga, 2011). For example, comparisons of the genomes of the Irish potato famine pathogen Phytophthora infestans and a closely related species Phytophthora mirabilis, which infects Mirabilis jalapa revealed that 82 of 345 genes which showed signs of positive selection could encode effector sequences (Raffaele et al., 2010). Subsequent work on orthologous effectors that encode protease inhibitor from the two species showed that the protease inhibitor effectors from each of these species interact specifically with protease targets from their respective host plants (Dong et al., 2014). Host-pathogen co-evolution is therefore reflected in adaptations of effectors for function in the host.

Heterodera sacchari is an increasingly economically important pathogen of several monocot species including rice and sugarcane. Crop losses due to H. sacchari can exceed 40%, with more severe damage reported under rain fed upland conditions (Kyndt et al., 2014). Compared to other characterised cyst nematodes, the biology of H. sacchari is unusual in that it is restricted to monocots and reproduces mainly by mitotic parthenogenesis (CABI, 2014). Here we sequenced and assembled the transcriptome of H. sacchari and use these data to reconstruct the evolutionary history of this and related species. These data suggest that H. sacchari and a related monocot parasite H. avenae evolved to parasitize monocot plants secondarily, from a dicot-parasitic ancestor. To explore the genetic changes associated with the evolutionary transition from dicot- to monocot-parasite, we identified homologues of previously identified effectors in H. sacchari. We show that in general the effectors have diversified in sequence more than non-effectors when compared to their most similar homologue in a dicot-parasite, and that while some effectors show conserved expression profiles and likely function, specific aspects of the effector repertoire of H. sacchari appear to be adapted to monocots.

3.3 Materials and Methods Biological material

Heterodera sacchari was cultured on rice cv Nipponbare as described in Pokhare et al. (2018). Briefly, plants were grown in a potting mixture of sand, field soil and organic matter (70:29:1) and were infected with second stage juveniles. After 12 weeks, watering of the

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari plants was stopped, and the plants were allowed to dry for 2 weeks. Cysts were collected by Cobb’s decanting and sieving method using standard protocols (Cobb, 1918). The cysts were surface sterilized and placed in 3mM ZnCl2 to initiate hatching. The resulting J2s were collected every 5 days and either frozen in liquid nitrogen or used immediately for infection of new plants grown on pluronic gel (Pokhare et al., 2018). Parasitic stage female nematodes were collected by hand 15 days after infection and frozen in liquid nitrogen until use.

Transcriptome sequencing

RNA was extracted from second stage juvenile and parasitic stage female nematodes using a Nucleospin RNA XS kit (Macherey Nagel) following the manufacturer’s instructions. The quantity and integrity of RNA were assessed using a Bioanalyzer. Library preparation for RNAseq was performed using the TruSeq RNA Library Prep Kit v2 (Illumina) as recommended by the manufacturer (Illumina; Protocol # 15026495, revision F). Separate libraries were generated from the juvenile (1µg) and female (500ng) total RNA samples, using single-end TruSeq Index Adapters AR002 (CGATGT) and AR004 (TGACCA), respectively. Each library was quality checked using a Bioanalyzer 2100 (Agilent) and quantified using a Qubit fluorometer (Thermo Fisher). Equal molarities of each library were combined and run at 12 pM on a MiSeq (Illumina) generating paired-end 2x 250 bp reads, as recommended. A fastq file was generated for each sample using MiSeq Control Software (version 2.6) for downstream QC and analysis.

Transcriptome assembly, quality control, and annotation

Scripts used to analyse the data are available at: https://github.com/peterthorpe5/Hsac_transcriptome. Assemblies are available at https://zenodo.org/deposit/1324265. Raw reads are available under primary accession PRJEB28025 and secondary accession ERP110186.

The 17725370 read pairs (5,636,376 from juveniles and 11,329,919 from females with 759075 undetermined) were first quality control checked using FastQC (Andrews, 2010), then quality trimmed using Trimmomatic version 0.32 (Q15) (Bolger et al., 2014). The resulting 17086132 read pairs were assembled using Trinity version 2.1.1 (kmer length 25) (Haas et al., 2013). The resulting assembly was subjected to quality control filtering using Transrate version 1.0.1 (Smith-Unna et al., 2016). Low quality/scoring transcripts were removed (based on read mapping to the assembly). Coding sequences were predicted using TransDecoder (Using DIAMOND BLASTP versus Swiss prot and HMM search versus Pfam

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

A domain guides). The predicted coding sequences were DIAMOND BLASTP version 0.7.9 (Buchfink et al., 2014) searched against GenBank NR database (May 2017). The results were interrogated for their alien index (AI) score using https://github.com/peterthorpe5/public_scripts/tree/master/Lateral_gene_transfer_prediction_ tool, which predicts putative horizontal gene transfer events. Any sequence with an AI greater than 25 and that had a BLAST hit identity greater than 70% to a non-metazoan was flagged as putative contamination. Putative contaminant sequences were removed, and the corresponding transcripts were removed from the assembly. BUSCO version 1.1b (Simão et al., 2015) and CEGMA version 2.4 (Parra et al., 2007) were used to quantify the completeness of the assembly. The resulting coding sequences were annotated using Trinotate (Grabherr et al., 2011), HMMER (Finn et al., 2011), Pfam (Yang & Smith 2013), SignalP 4.0 (Petersen et al., 2011), TMHMM (Krogh et al., 2001), BLAST (Altschul et al., 1990), gene ontology (Ashburner et al., 2000), eggNOG V3.0 (Powell et al., 2012) and RNAmmer (Lagesen et al., 2007). SPRYSECs were identified using HMMER, and Phobius (Käll et al., 2007) for those containing a secretion signal. CAZYme analysis was performed by the CAYZme analysis toolkit using both BLAST and PFAM analysis on the online server (Park et al., 2010). BLAST2GO was used to identify and analyse GO terms (Conesa et al., 2005). The H. sacchari transcripts were compared to all genes predicted in the G. pallida and G. rostochiensis genomes with matches identified using an e value cut off of 10-10. Similarly, candidate H. sacchari effectors were identified by comparisons with effectors from other cyst nematode species (Gao et al., 2003; Thorpe et al., 2014; Eves-van den Akker et al., 2016a) using an e value cut off of 10-10.

Phylogenetics

One hundred and one CEGMA genes conserved in the genome and or transcriptome resources of 18 plant-parasitic nematodes species were used for phylogenetic reconstruction. The species used were H. avenae (Kumar et al., 2014), H. glycines (T. Baum, pers. comm.), G. pallida (Cotton et al., 2014), G. rostochiensis (Eves-van den Akker et al., 2016a), G. ellingtonae (Phillips et al., 2017), R. reniformis (Eves-van den Akker et al., 2016b), R. similis (Jacob et al., 2008), M. arenaria (Blanc-Mathieu et al., 2017), M. javanica (Blanc-Mathieu et al., 2017), M. incognita (Blanc-Mathieu et al., 2017), M. hapla (Opperman et al., 2008), P. coffeae (Burke et al., 2015), N. aberrans (Eves-van den Akker et al., 2014), B. xylophilus (Kikuchi et al., 2011), C. elegans (C. elegans sequencing consortium, 2018), C. briggsae (Hillier et al., 2007) and L. elongatus (Danchin et al., 2017). The protein sequences of

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

CEGMA genes were aligned and refined individually using MUSCLE (Edgar, 2004). Individual alignments were concatenated and submitted to the IQtree online server with associated partition file. Model selection was carried out on each partition and a concatenated multi-gene phylogeny generated using the ultra-fast mode and 1000 bootstraps (Trifinopoulos et al., 2016).

CLE effector identification and network analysis

Several partial transcripts encoding proteins with similarity to CLE-effectors from various plant-parasitic nematodes were identified in the transcriptome assembly. Partial transcripts were computationally extended using an iterative approach of mapping and overlap assembly using the wrapper script provided with MITOBIM (Hahn et al., 2013). Only the deduced amino acid sequences of full length CLEs were used for further analyses. To compare H. sacchari CLE domains to those of plants a database of CLE peptides was collated from Zhang et al. (2014), and Oelkers et al. (2008). All CLE domains were combined into a fasta file and filtered for those with missing information. Custom python script 1 was used to construct an all vs all similarity matrix based on the BLOSUM62 scores between amino acids. Custom python script 2 was used to parse the matrix and normalise self-normalise similarity scores. Custom python script 3 was used to convert the matrix to gefx, which was loaded into Gephi to visualise the network (Bastian et al., 2009). Custom python scripts are available under github repository https://github.com/sebastianevda/.

Cloning and characterisation of candidate effector sequences

The complete ORFs of selected genes were amplified from cDNA of pre-parasitic second stage juveniles. PCR products were purified using the Qiagen PCR Purification kit and cloned into the pGEMT Easy or pCR8/TOPO/GW vectors following the manufactures guidelines.

Analysis expression profiles of candidate effectors

The spatial expression patterns of candidate effectors from H. sacchari were investigated using in situ hybridisation of digoxigenin labelled probes to juvenile nematodes as previously described (Thorpe et al., 2014). The expression of candidate effectors across the H. sacchari life cycle (eggs, pre-parasitic juveniles, 15 & 25 days post infective females) was analysed using quantitative PCR (qRT-PCR) with gene specific primers. Around 3000 eggs and pre- parasitic juveniles were collected for each replicate. To collect eggs, cysts were crushed under a binocular microscope and eggs were transferred into an Eppendorf tube in sterile

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari water. Juveniles were harvested every 5 days as described above and flash frozen in liquid nitrogen. For the other life stages (15 & 25 days post inoculation), approximately 150 parasitic stage nematodes were collected from rice grown in pluronic gel as described above. Nematodes were stored at -80° C and used subsequently for RNA extraction. Total RNA was extracted as above and were processed using Agilent 2100 bioanalyzer system to check the quality and quantity of extracted RNA. The RNA samples with RNA integrity number (RIN) of more than 8 were used for cDNA synthesis using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems) and oligo-dT primer. The cDNA was tested for expression changes during the nematode life stages using the Stepone Plus Real-Time PCR System (Applied Biosystems) with cycling parameters of 95° C for 15 s and 60°C for 30 s (40 cycles) for amplification. Each sample well contained 10 μl of Fast SYBR Green qPCR Master Mix (Invitrogen), 9 μl of the gene specific primer mixture with a final concentration 1 μM for each primer and 1 μl of cDNA. The data was analysed using the Stepone Plus Real- Time PCR software to create Ct values and relative expression was calculated following Pfaffl (2001). Elongation Factor 1 alpha (EF1 alpha) was used as internal control for all experiments. Three biological replicates from each stage and three technical replicates for each biological replicate were used for qPCR studies.

Functional analysis of H. sacchari CLE sequences

To analyse the in vivo function of CLE peptides, we developed a protocol for exogenous application to rice seedlings, similar to that described for A. thaliana (Wang et al., 2010). The terminal 13 amino acid CLE domain of nematode CLE-like sequences was synthesised with hydroxy prolines in positions 5 and 8 for H. sacchari (H-Lys-Arg-Leu-Ser-Hyp-Gly-Gly- Hyp-Asp-Pro-Gln-His-His-OH), H. glycines (H-Lys-Arg-Leu-Ser-Hyp-Ser-Gly-Hyp-Asp- Pro-His-His-His-OH). As a control, a shuffled version of the H. sacchari CLE sequence was also synthesised (H-Asp-His-Ser-Hyp-Gly-Gln-His-Arg-Hyp-Lys-Pro-Gly-Leu-OH). Seeds of rice (cv Nipponbare) were surface sterilised and allowed to germinate on sterile, wet filter paper. After 7 days, plants of similar size were transferred to plates containing ½ MS supplemented with 10 µM of the relevant peptide or an equivalent volume of sterile distilled water and left for 10 days at 25 oC in 16h light/8h dark. After this time plants were removed from the plates and the roots were washed to remove any adhering medium. The length of the longest root was measured for 24 biological replicates of each condition.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

3.4 Results and Discussion

The transcriptome of H. sacchari

A total of 17,086,132 paired end reads were obtained from the cDNA extracted from second stage juveniles (J2) and parasitic stage nematodes 15 days after infection. These sequence reads have been submitted to SRA (accession number PRJEB28025). The sequence reads were pooled and assembled into a single reference transcriptome of 44,230 transcripts after filtering with Transrate and removing contaminants. Assemblies are available at https://zenodo.org/deposit/1324265. CEGMA analysis showed that 88% of the core eukaryotic genes were present as full-length transcripts, with a further 6% represented by partial length transcripts. BUSCO analysis using the metazoan data set suggests that the assembly contains 76% complete BUSCO sequences and a further 6.0% fragmented sequences. Seventeen percent of the BUSCO genes were not identified. Comparisons with published gene models from cyst nematode genome sequences, showed that 34,203 (77.3%) and 30,482 (68.9%) of the H. sacchari transcripts matched sequences in G. rostochiensis and G. pallida respectively (evalue 1e-10). Taken together, these data suggest that the assembly produced represents a large proportion of H. sacchari transcriptome of these life stages.

H. sacchari evolved to parasitise monocotyledonous plants from an ancestor parasitic on dicotylendonous plants

We used a subset of 101 core eukaryotic genes conserved in H. sacchari and 17 related species to reconstruct a multi-gene phylogeny (Figure 1). This phylogeny robustly positions H. sacchari and the related cereal cyst nematode H. avenae in a mono-phyletic sub-clade of monocot parasites, nested within a clade of related dicot-parasites of the genera Heterodera and Globodera. The most parsimonious explanation for this is that 1) H. sacchari and H. avenae share a common, monocot-parasitic, ancestor, and 2) this last common monocot- parasitic ancestor evolved to parasitise a monocot host secondarily, from a dicot-parasitic ancestor. This provides the comparative framework to explore the genes conserved, and the genes diverged, during adaptation to monocot parasitism by nematodes.

Genes encoding cell wall modifying enzymes acquired via horizontal gene transfer in the H. sacchari transcriptome

The plant cell wall is the first significant barrier to an invading pathogen, and while largely similar between dicots and monocot, there are noticeable differences in composition. Plant- parasitic nematodes in general are well equipped with proteins that allow them to modify and

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari degrade specific components of the plant cell wall. Many of these genes were acquired via horizontal gene transfer from bacteria (Danchin et al., 2010).

The transcriptome of H. sacchari contains representatives of most previously described cases of horizontal gene transfer in related plant-parasitic nematodes, including a wide range of cell wall degrading enzymes (Table 1) and several other sequences putatively acquired by horizontal gene transfer (e.g. the GH32 invertases and chorismate mutase (Danchin et al., 2016; Jones et al., 2003)). Analysis of the expression profiles of one of the H. sacchari GHF5 cellulases and the chorismate mutase showed that, as in other cyst nematodes, expression was restricted to the subventral pharyngeal gland cells in J2s and while the cellulase was upregulated at J2 the chorismate mutase was expressed throughout the life cycle (Figure 2). These sequences may therefore play a similar role in the biology of H. sacchari and other cyst nematodes.

Notably however, both the H. sacchari and H. avenae transcriptomes lack sequences similar to GH53 Arabinogalactan endo- 1,4 beta-galactosidase, in spite of the fact that all other cyst nematodes analysed to date, which parasitise dicots, have such proteins. While the absence of evidence in transcriptome datasets is not necessarily evidence of absence, it nevertheless reflects the host range of these species. Cell walls of commenlinoid monocots (which include the main hosts of H. sacchari and H. avenae) have a different composition compared to those of dicots and contain relatively low amounts of pectic polysaccharides, including the substrate of the GH53 enzymes (Vogel, 2008). Without genomic resources we cannot conclude whether the conspicuous absence in the transcriptomes of the monocot parasites is because these genes have been lost entirely or that they are not expressed under these conditions.

An overview of H. sacchari effector-like sequences

Effectors modulate plant process to promote disease and are often finely tuned to their host. To determine whether the effector repertoire of H. sacchari reflects its secondary adaptation to a monocot host, we first identified and characterised effectors in the H. sacchari transcriptome by building on a detailed genome wide analysis of cyst nematode effectors performed for G. rostochiensis (Eves-van den Akker et al., 2016a). Using G. rostochiensis effectors as a starting point for comparative purposes, 185 of the 295 identified a similar sequence in the H. sacchari transcriptome (at e value <1e-10). We compared the similarity of H. sacchari effector-like sequences and non-effector-like sequences to their most similar

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari homologue in the G. rostochiensis genome. This showed that, on average, putative effectors of H. sacchari are more different to their closest homologue in G. rostochiensis than non- effectors are to their corresponding closest homologue (Figure 3). Taken together, these data suggest that while most effectors are apparently conserved, they have nevertheless accumulated more mutations that than non-effectors since these species diverged.

Consistent with this observation, analysis of effectors from a range of pathogens, including PPN, has shown that they are under strong diversifying selection pressure. For example, whole genome resequencing of five pathotypes of G. rostochiensis showed that effectors contain significantly more variants and more nonsynonymous variants per gene than do randomly selected non-effector genes (Eves-van den Akker et al., 2016a). On a more detailed scale, the SPRYSEC effector (RBP1) from G. pallida, which is recognised by the Gpa2 resistance gene of potato, has been subjected to positive selection at several different residues, including the residue that determines recognition or evasion by Gpa2 (Sacco et al., 2009).

The SPRY domain family is greatly expanded in cyst nematodes with 295 sequences predicted in G. pallida of which approximately 30 have a signal peptide (and thus encode SPRYSEC effectors) (Cotton et al., 2014), and 71 sequences in G. rostochiensis of which 17 may be SPRYSEC effectors (Eves-van den Akker et al., 2016a). While we find a similar proportion of SPRYs to SPRYSECs in H. sacchari (80 SPRY domain encoding transcripts and 6 SPRYSECs), with similar spatial expression in the dorsal pharyngeal gland cell (Figure 4), the temporal expression is unusual. At least two H. sacchari SPRYSECs are upregulated in parasitic stage nematodes (Figure 4) and this contrasts with the G. pallida SPRYSECs which tend to be upregulated in J2s (Mei et al., 2015). Several SPRYSECs have been shown to suppress host defence signalling (Postma et al., 2012; Mei et al., 2015; Mei et al., 2018). What role the H. sacchari SPRYSECs play in infection is yet to be determined.

Other notable peculiarities in the effector repertoire of H. sacchari are the HYP effectors. The HYP effectors, first identified in G. pallida and G. rostochiensis, are secreted from the gland cells surrounding the main anterior sense organs and show unprecedented variability between individuals (Eves-van den Akker et al., 2014). The HYP effectors are strongly upregulated in parasitic stages of G. pallida and can be subdivided into three subfamilies based on the presence and type of subfamily-specific tandem repeats. The transcriptome of H. sacchari contains two full length HYP-like transcripts (primarily represented in the 15 days post infection library) with unusual characteristics. A phylogenetic analysis of the H. sacchari

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

HYP-like sequences with all other known full-length HYP effectors (n = 65) shows that the H. sacchari HYP-like sequences group in a separate subfamily (named Subfamily HYP-0, Figure 5). Comparison of the H. sacchari predicted sequences with those from G. pallida shows that these sequences have both highly conserved regions that flank the variable domain but they are the first to encode a major structural variant outside the hypervariable domain, and the region corresponding to the “hypervariable domain” contains a novel sequence that has a very limited repeat structure (sequence features summarised in Figure 5).

Specialisation of H. sacchari CLE-like effectors to parasitism of a monocotyledonous host

One class of H. sacchari effectors shows signs of adaptation to a monocot host. The CLAVATA3/EMBRYO SURROUNDING REGION-like (CLE) effectors mimic plant peptide hormones and have been characterised in a number of plant-parasitic nematodes. The transcriptome of H. sacchari contained several partial transcripts that encode proteins with similarity to CLE-like effectors from other plant-parasitic nematodes. We were able to computationally extend seven unique transcripts to recapitulate the full-length open reading frame, resulting in 5 unique polypeptide sequences from methionine to stop. As for other cyst nematodes, the H. sacchari sequences each encoded a signal peptide at the N-terminus followed by an N-terminal domain (Figure S1), which in other cyst nematodes enables translocation into the apoplast after the protein is secreted into the plant cell (Wang et al., 2010a). Six of these H. sacchari transcripts also encode a single canonical CLE domain at their C terminus, while the seventh encodes a tandemly repeated motif with no clear homology to canonical CLE domains (despite the similarity of the rest of the protein sequence to CLE-effectors of other cyst nematodes). The six H. sacchari CLE-effector-like sequences can be divided into two groups based at least in part on their signal peptide and CLE domains. The CLE domains within each group are identical in protein and nucleic acid sequence (Figure 6A).

Given that CLE peptides vary between plant species in general, and monocots and dicots in particular, we hypothesised that the CLE domains of CLE-like effectors in H. sacchari may have specialised prior to/concurrent with the transition from dicot- to monocot-parasite. To test this hypothesis, we analysed a database of 391 CLE peptides from 20 plant species collated from Zhang et al. (2014) and Oelkers et al. (2008). We created an all by all matrix of similarity between plant CLE peptides and H. sacchari CLE peptides based on a normalized BLOSUM62 score. We then used this matrix to generate a CLE similarity network (Figure

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

6B), highlighting the host (rice Oryza sativa) and the nematode (H. sacchari) CLEs. Three of the H. sacchari CLEs form part of a well-connected cluster containing 14 other CLEs, 8 of which are from rice. On closer inspection, we found that three of these H. sacchari CLEs (Hsac_DN37996_c0_g4_i1, Hsac_DN35920_c0_g3_i1 and Hsac_DN35920_c0_g2_i1) were sequence-identical mimics of eight rice CLEs (OS_GEN_Os01g55080_1, OSEST_NP890021__1, OSEST_TC269510__1, OS_TA_AK108976__1, OS_TA_CA758496__1, OS_GEN_Os01g48230_1, OS_GEN_Os01g48260_1 and OSEST_TC271220__1) in the terminal 13 amino acids of its CLE domain. Three other H. sacchari CLEs (Hsac_DN37996_c0_g1_i1, Hsac_DN49341_c0_g1_i1 and Hsac_DN35920_c0_g1_i1) form a small cluster with one other rice CLE (OS_GEN_Os05g48730_1). The terminal 13 amino acids of the remaining CLE-effector-like sequence that lacks a canonical CLE domain has no connections in the network. H. sacchari CLEs are the only nematode CLEs with a connection to a rice CLE. Interestingly, there are several other sequence divergent CLE clusters in rice with no corresponding H. sacchari CLE, suggesting selective mimicry of a subset of this family. It is known that CLE family members in dicots have diverse roles (e.g. Mitchum et al., 2008) and mimicry of a subset of rice CLEs by H. sacchari may reflect the need to target the function of a subset of the full rice CLE complement.

The functional significance of the similarity between the H. sacchari CLE peptides and those from rice was experimentally investigated by taking advantage of the short root phenotype observed from overexpression or exogenous application of CLEs (e.g. Fiers et al., 2004 and Chen et al., 2015). We synthesised a synthetic version of the H. sacchari CLE peptide that had the highest connectivity with rice CLEs in the network (sequence identical to the rice CLEs in the 13 terminal amino acids), and exogenously applied it to rice seedlings. We then analysed the effect on root growth when compared to a randomised version of this CLE, or a CLE from the dicot-parasitic H. glycines. This analysis showed that the peptide from H. sacchari induced a short root phenotype in rice whereas peptides from H. glycines and a shuffled peptide used as a control had no effect (n=24 per condition, p <0.001, Tukey’s HSD, Figure 7).

3.5 Conclusions

We use whole transcriptome sequencing to show that H. sacchari and the related H. avenae evolved to parasitise a monocot host from a last common dicot-parasitic ancestor. We mine these data to identify and characterise the cell wall degrading enzyme and effector

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari complement of H. sacchari. Finally, we show that while H. sacchari has a similar effector arsenal to the related cyst nematodes that parasitise dicots, the CLE effectors provide a paradigm of functional adaptation for parasitism of a monocot host. These data build a foundation on which to explore novel effectors involved in monocot-parasitism.

Acknowledgements

Somnath Pokhare was supported by a fellowship provided by the Indian Council of Agricultural Research (ICAR), Government of India. This work was also supported by Royal Society International Exchanges award number IE150670 and by the Rural and Environmental Science and Analytical Services Division of the Scottish Government. SEvdA is supported by Biotechnology and Biological Sciences Research Council grant BB/R011311/1. This work benefited from interactions funded through COST Action FA1208.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

3.6 Tables Table 3.1: H. sacchari sequences similar to putative cell wall degrading and modifying proteins

Substrate Cellulose Xylan Arabi Pectin Arabino nan galactan Family GH5 GH45 GH30 GH43 GH PL GH53 Expan CBM2 28 3 sins domains H. sacchari 8 0 0 0 0 1 0 2 2 H. avenae 16 0 0 0 0 2 0 2 2 H. schachtii 9 0 1 0 9 3 4 2 G. rostochiensis 11 0 0 1 0 3 1 7 7 G. pallida 16 0 0 1 0 7 2 9 6 M. incognita 21 0 6 2 1 30 0 20 9 N. aberrans 2 0 3 1 2 8 3 2 2 B. xylophilus 0 11 0 0 0 15 0 8 0

Table 3.S1: Primers used For In situ hybridisation

Hsac_Cellulase F CCTTGGCACTCCGATGTATT Hsac_Cellulase R ATCAGCTCGTCCAGCATTTT Hsac_Chorismate mutase F GCTGCTTTAACTTCCGCAAC Hsac_Chorismate mutase R CCGTTTCTTGGTTCGGTTTA Hsac_DN36333 F TGTCAAAGGAGCGAGGGATT Hsac_DN36333 R ATGAGGCTTCCACAGTTCCA Hsac_DN43534 F AAATTCCCAGCTGCGATCAC Hsac_DN43534 R GTCGGAAGCAAATCGGTCTC

Table 3.S2: Primers used for quantitative PCR (qRT-PCR)

Hsac_Cellulase qF CTGGACCGACGAAACAGTGA Hsac_Cellulase qR TCGATCGCTTTGTCCGGAAA Hsac_Chorismate mutase qF AGCATTGACGACTTTGTGCG Hsac_Chorismate mutase qR ATTTCGTCGGTTGCCTCTGT Hsac_DN36333 qF GGGGCCATTCTGTCAAAGGA Hsac_DN36333 qR TGAGGCTTCCACAGTTCCAC Hsac_DN43534 qF AGAAACGTCGACAGTGGCAT Hsac_DN43534 qR GACAAGCGCAAACTCAGCAA

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

3.7 Figures

Figure 3.1: Phylogenetic analysis of 101 CEGMA genes conserved in 18 species of nematode shows that H. sacchari and H. avenae share a common monocot parasitic ancestor that most likely evolved from a parasite of dicots.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Figure 3.2: Analysis of expression patterns of cellulase (left panels) and chorismate mutase (right panels) identified in the H. sacchari transcriptome. In situ hybridisation analysis showed that both genes are expressed in the subventral gland cells of second stage juveniles (J2s) of H. sacchari (purple staining; upper panels). Analysis using qPCR shows that cellulase is expressed specifically at the J2 life stage while the chorismate mutase is expressed at all life stages tested (lower panels).

Figure 3.3: Histograms showing distributions of percentage identity between H. sacchari effectors (left panel) and non-effectors (right panel) with their homologues in G. rostochiensis. This analysis suggests that effector-like transcripts have accumulated more mutations than other transcripts since the divergence of these two species.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Figure 3.4: Analysis of expression patterns of two putative SPRYSEC-encoding transcripts of H. sacchari. In situ hybridisation analysis showed that both genes are expressed in the dorsal gland cell of H. sacchari (purple staining, arrows; upper panels). Analysis by qPCR shows that both sequences are upregulated in later parasitic stage nematodes (lower panels).

Figure 3.5: Overview of the HYP gene family in cyst nematodes. A schematic representation of the H. sacchari HYP-like sequences shows the position of a novel structural variant outside the hyper-variable domain. A midpoint re-rooted phylogenetic construction of HYP sequences from G. pallida and H. sacchari position those of H. sacchari as a distinct subfamily (names here sub-family-0).

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Figure 3.6: Alignment of CLE-effector-like sequences of H. sacchari. Highlighted the position of the predicted signal peptide, the unknown repeat of DN37996_c0_g2_i1, and the canonical CLE domain of the other six sequences.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Figure 3.7: CLE-like effectors of H. sacchari. A) A midpoint re-rooted phylogenetic construction of the seven CLE-like sequences in the H. sacchari transcriptome. Six of these sequences encode a canonical CLE domain at the C terminus, while the seventh encodes tandem repeats of an unknown peptide. The CLE domain containing effectors form two groups, each of which has a distinct CLE domain. B) A similarity network of CLE domains from plants and H. sacchari. Rice CLEs are highlighted in green, and H. sacchari CLE-like sequences in orange. Nodes in the network are scaled by connectivity.

Figure 3.8: Functional adaptation of H. sacchari CLE effectors to monocot hosts. Exogenous application of synthetic H. sacchari CLE peptides causes a short root phenotype when compared to a scrambled version of this peptide, a CLE peptide from the related dicot parasitic H. glycines, or no peptide control (n=24 per condition, p <0.001, Tukey’s HSD).

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

3.8 References Altschul, S.F., Gish, W., Miller, W., Myers, E.W. and Lipman, D.J. (1990). Basic local alignment search tool. Journal of Molecular Biology 215(3), 403-410.

Andrews S. (2010). FastQC: a quality control tool for high throughput sequence data. Available online at: http://www.bioinformatics.babraham.ac.uk/projects/fastqc.

Ashburner, M., Ball, C.A., Blake, J.A., Botstein, D., Butler, H., Cherry, J.M., Davis, A.P., Dolinski, K., Dwight, S.S., Eppig, J.T. and Harris, M.A. (2000). Gene Ontology: tool for the unification of biology. Nature Genetics 25(1), 25-29.

Bastian M., Heymann S. and Jacomy M. (2009). Gephi: an open source software for exploring and manipulating networks. International AAAI Conference on Weblogs and Social Media.

Blanc-Mathieu, R., Perfus-Barbeoch, L., Aury, J.M., Da Rocha, M., Gouzy, J., Sallet, E., et al. (2017). Hybridization and polyploidy enable genomic plasticity without sex in the most devastating plant-parasitic nematodes. PLoS genetics 13(6), 1-36.

Bolger, A.M., Lohse, M., and Usadel, B. (2014). Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics 30(15), 2114-2120.

Buchfink, B., Xie, C. and Huson, D.H. (2015). Fast and sensitive protein alignment using DIAMOND. Nature Methods 12, 59-60.

Burke, M., Scholl, E.H., Bird, D. McK., Schaff, J.E., Colman, S.D., Crowell, R., Diener, S., Gordon, O., Graham, S., Wang, X., Windham, E., Wright, G.M. and Opperman, C.H. (2015). The plant parasite Pratylenchus coffeae carries a minimal nematode genome. Nematology 17(6), 621-637.

C. elegans sequencing consortium (1998). Genome sequence of the nematode C. elegans: A platform for investigating biology. Science 282, 2012-2018.

CABI (2014). Heterodera sacchari. 139, 43–45.

Chen, S., Lang, P., Chronis, D., Zhang, S., De Jong, W.S., Mitchum, M.G. and Wang, X. (2015). In planta processing and glycosylation of a nematode CLAVATA3/ENDOSPERM SURROUNDING REGION-like effector and its interaction with a host CLAVATA2-like receptor to promote parasitism. Plant Physiology 167(1), 262-272.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Chronis, D., Chen, S., Lu, S., Hewezi, T., Carpenter, S.C.D., Loria, R., Baum, T.J. and Wang, X. (2013). A ubiquitin carboxyl extension protein secreted from a plant-parasitic nematode Globodera rostochiensis is cleaved in planta to promote plant parasitism. The Plant Journal 74(2), 185-196.

Cobb, N.A. (1918). Estimating the nema population of soil, with special reference to the sugar-beet and root-gall nemas, Heterodera schachtii Schmidt and Heterodera radicicola (Greef) Müller and with a description of Tylencholaimus aequalis n. sp. Washington, DC, USA, National Government Printing Office.

Conesa, A., Götz, S., García-Gómez, J.M., Terol, J., Talón, M. and Robles, M. (2005). Blast2GO: a universal tool for annotation, visualization and analysis in functional genomics research. Bioinformatics 21(18), 3674-3676.

Craig, J.P., Bekal, S., Hudson, M., Domier, L., Niblack, T. and Lambert, K.N. (2008). Analysis of a horizontally transferred pathway involved in vitamin B6 biosynthesis from the soybean cyst nematode Heterodera glycines. Molecular Biology and Evolution 25(10), 2085-2098.

Danchin E.G.J., Rosso, M.N., Vieira. P., de Almeida-Engler, J., Coutinho, P.M., Henrissat, B. and Abad, P. (2010). Multiple lateral gene transfers and duplications have promoted plant parasitism ability in nematodes. Proceedings of the National Academy of Sciences 107(41), 17651–17656.

Danchin, E.G.J., Guzeeva, E., Mantelin, S., Berepiki, A. and Jones, J.T. (2016). Horizontal gene transfer from bacteria has enabled the plant-parasitic nematode Globodera pallida to feed on host-derived sucrose. Molecular Biology and Evolution 33(6), 1571-1579.

Danchin, E.G.J., Perfus-Barbeoch, L., Rancurel, C., Thorpe, P., Da Rocha, M., Bajew, S., Neilson, R., Sokolova, E., Da Silva, C., Guy, J., Labadie, K., Esmenjaud, D., Helder, J., Jones, J.T. and Eves-van den Akker, S. (2017). The Transcriptomes of Xiphinema index

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

and Longidorus elongatus suggest independent acquisition of some plant parasitism genes by horizontal gene transfer in early-branching nematodes. Genes 8(10), 287.

Dong, A., Stam, R., Cano, L.M., Song, J., Sklenar, J., Yoshida, K., Bozkurt, T.O., Oliva, R., Liu, Z., Tian, M., Win, J., Banfield, M.J., Jones, A.M.E., van der Hoorn, R.A.L. and Kamoun, S. (2014). Effector specialization in a lineage of the Irish Potato Famine Pathogen. Science 343(6170), 552-555.

Edgar, R.C. (2004). MUSCLE: a multiple sequence alignment method with reduced time and space complexity. BMC Bioinformatics 5:113.

Espada, M., Silva, A.C., Eves-van den Akker, S., Cock, P.J.A., Mota, M. and Jones, J.T. (2016). Identification and characterization of parasitism genes from the pine wilt nematode Bursaphelenchus xylophilus reveals a multi-layered detoxification strategy. Molecular Plant Pathology 17(2), 286-295.

Espada, M., Eves-van den Akker, S., Maier, T., Vijaypalani, P., Baum, T.J., Mota, M. and Jones, J.T. (2018). STATAWAARS: A promoter motif associated with the tissue- specific transcriptome of the major effector-producing glands of the plant-parasitic nematode Bursaphelenchus xylophilus. BMC Genomics 19: 553.

Eves-van den Akker, S., Lilley, C.J., Jones, J.T. and Urwin, P.E. (2014). Identification and characterisation of a hyper variable apoplastic effector gene family of G. pallida. PLoS Pathogens 10(9): e1004391.

Eves-van den Akker, S. and Birch, P.R.J (2016). Opening the effector protein toolbox for plant–parasitic cyst nematode interactions. Molecular Plant 9(11), 1451-1453.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

of the yellow potato cyst nematode, Globodera rostochiensis, reveals insights into the bases of parasitism and virulence. Genome Biology 17, 124.

Eves-van den Akker, S., Lilley, C.J., Yusup, H.B., Jones, J.T. and Urwin, P.E. (2016b). Functional C-terminally encoded plant peptide (CEP) hormone domains evolved de novo in the plant parasite Rotylenchulus reniformis. Molecular Plant pathology 17(8), 1265–1275.

Fiers, M.A., Hause, G., Boutilier, K.A., Casamitjana-Martinez, E., Weijers, D., Offringa, R., van der Geest, L., Lookeren Campagne, M.van and Liu, C.M. (2004). Mis-expression of the CLV3/ESR-like gene CLE19 in Arabidopsis leads to a consumption of root meristem. Gene, 327(1), 37– 49.

Finn, R.D., Clements, J. and Eddy, S.R. (2011). HMMER web server: interactive sequence similarity searching. Nucleic Acids Research 39, W29-W37.

Gao, B., Allen, R., Maier, T., Davis, E.L., Baum, T.J. and Hussey, R.S. (2003). The parasitome of the phytonematode Heterodera glycines. Molecular Plant-Microbe Interactions 16(8), 720–726.

Gheysen, G. and Mitchum, M.G. (2011). How nematodes manipulate plant development pathways for infection. Current Opinion in Plant Biology 14(4), 415-421.

Grabherr, M.G., Haas, B.J., Yassour, M., Levin, J.Z., Thompson, D.A., Amit, I., Adiconis, X., Fan, L., Raychowdhury, R. and Zeng, Q. (2011). Full-length transcriptome assembly from RNA-Seq data without a reference genome. Nature Biotechnology 29(7), 644-652.

Haas, B.J., Papanicolaou, A., Yassour, M., Grabherr, M., Blood, P.D., Bowden, J., Couger, M.B., Eccles, D., Li, B., Lieber, M., MacManes, M.D., Ott M., Orvis, J., Pochet, N., Strozzi, F., Weeks, N., Westerman, R., William, T., Dewey, C.N., Henschel, R., LeDuc, R.D., Friedman, N. & Regev, A. (2013). De novo transcript sequence reconstruction from RNA-seq using the Trinity platform for reference generation and analysis. Nature Protocols 8, 1494-1512.

Hahn, C., Bachmann, L., and Chevreux, B. (2013). Reconstructing mitochondrial genomes directly from genomic next-generation sequencing reads—a baiting and iterative mapping approach. Nucleic acids research 41(13), e129.

Hewezi, T., Howe, P., Maier, T., Hussey, R.S., Mitchum, M.G., Davis, E.L. and Baum, T.J. (2008). Cellulose binding protein from the parasitic nematode Heterodera schachtii

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

interacts with Arabidopsis pectin methylesterase: cooperative cell wall modification during parasitism. Plant Cell 20(11), 3080-3093.

Jacob, J., Mitreva, M., Vanholme, B. and Gheysen, G. (2008). Exploring the transcriptome of the burrowing nematode Radopholus similis. Molecular Genetics and Genomics 280(1), 1-17.

Hillier, L.W., Miller, R.D., Baird, S.E., Chinwalla, A., Fulton, L.A., Koboldt, D.C. and Waterston, R.H. (2007). Comparison of C. elegans and C. briggsae genome sequences reveals extensive conservation of chromosome organization and synteny. PLoS biology 5(7), e167.

Jones, J.T., Furlanetto, C., Bakker, E., Banks, B., Blok, V.C., Chen, Q. and Prior, A. (2003). Characterisation of a chorismate mutase from the potato cyst nematode Globodera pallida. Molecular Plant Pathology 4(1), 43-50.

Jones, J.T and Mitchum, M.G (2018). Biology of effectors. In Perry, R.N., Moens, M & Jones, J.T. (eds.). Cyst Nematodes. Wallingford, CABI, pp. 74-88.

Käll, L., Krogh, A. and Sonnhammer, E.L. (2007). Advantages of combined transmembrane topology and signal peptide prediction—the Phobius web server. Nucleic Acids Research 35 (suppl_2), W429-W432.

Kikuchi, T., Cotton, J.A., Dalzell, J.J., Hasegawa, K., Kanzaki, N., McVeigh, P., Takanashi, T., Tsai, I.J., Assefa, S.A., Cock, P.J.A., Otto, T.D., Hunt, M., Reid, A.J., Sanchez- Flores, A., Tsuchihara, K., Yokoi, T., Larsson, M.C., Miwa, J., Maule, A.G., Sahashi, N. Jones, J.T. and Berrima, M. (2011). Genomic insights into the origin of parasitism in the emerging plant pathogen Bursaphelenchus xylophilus. PLoS Pathogens 7(9), e1002219.

Kikuchi, T., Eves-van den Akker, S. and Jones, J.T. (2017). Genome evolution of plant- parasitic nematodes. Annual Review of Phytopathology 55, 333-354.

Krogh, A., Larsson, B., Von Heijne, G. and Sonnhammer, E.L. (2001). Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. Journal of Molecular Biology 305(3), 567-580.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Kyndt, T., Fernandez, D. and Gheysen G. (2014). Plant-parasitic nematode infections in rice: molecular and cellular insights. Annual Review of Phytopathology 52, 135-153.

Lagesen, K., Hallin, P., Rødland, E.A., Stærfeldt, H.H., Rognes, T. and Ussery, D.W. (2007). RNAmmer: consistent and rapid annotation of ribosomal RNA genes. Nucleic Acids Research, 35(9), 3100-3108.

Lee, C., Chronis, D., Kenning, C., Peret, B., Hewezi, T., Davis, E.L., Baum, T.J., Hussey, R.S., Bennet, M. and Mitchum, M.G. (2011). The novel cyst nematode effector protein 19C07 interacts with the Arabidopsis auxin influx transporter LAX3 to control feeding site development. Plant Physiology 155(2), 866-880.

Maier, T.R., Hewezi, T., Peng, J.Q. and Baum, T.J. (2013). Isolation of whole esophageal gland cells from plant-parasitic nematodes for transcriptome analyses and effector identification. Molecular Plant-Microbe Interactions 26(1), 31-35.

Mei, Y., Thorpe, P., Guzha, A., Haegeman, A., Blok, V.C., MacKenzie, K., Gheysen, G., Jones, J.T and Mantelin, S. (2015). Only a small subset of the SPRY domain gene family in Globodera pallida is likely to encode effectors, two of which suppress host defences induced by the potato resistance gene Gpa2. Nematology 17(4), 409-424.

Mei, Y., Wright, K.M., Haegeman, A., Bauters, L., Diaz-Granados, A., Goverse, A., Gheysen, G., Jones, J.T. and Mantelin, S. (2018). The Globodera pallida SPRYSEC effector GpSPRY-414-2 that suppresses plant defenses targets a regulatory component of the dynamic microtubule network. Frontiers in Plant Science 9:1019.

Mitchum, M.G., Wang, X. and Davis, E.L. (2008). Diverse and conserved roles of CLE peptides. Current Opinion in Plant Biology 11(1), 75-81.

Nicol J.M., Turner S.J., Coyne D.L., Nijs L.D., Hockland S., Maafi Z.T. (2011). Current nematode threats to world agriculture. In: Genomics and Molecular Genetics of Plant- Nematode Interactions (Jones J., Gheysen G., Fenoll C. eds.), pp. 21-43. Springer, Dordrecht.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Oelkers, K., Goffard, N., Weiller, G.F., Gresshoff, P.M., Mathesius, U. and Frickey, T. (2008). Bioinformatic analysis of the CLE signaling peptide family. BMC Plant Biology 8(1), 1.

Park, B.H., Karpinets, T.V., Syed, M.H., Leuze, M.R. and Uberbacher, E.C. (2010). CAZymes Analysis Toolkit (CAT): web service for searching and analyzing carbohydrate-active enzymes in a newly sequenced organism using CAZy database. Glycobiology 20(12), 1574-1584.

Parra, G., Bradnam, K. and Korf, I. (2007). CEGMA: a pipeline to accurately annotate core genes in eukaryotic genomes. Bioinformatics 23(9), 1061-1067.

Perry, R.N., Moens, M and Jones, J.T. (2018). Cyst Nematodes. Wallingford, CABI.

Popeijus, H., OIvermars, H., Jones, J.T., Blok, V.C., Goverse, A., Helder, J., Schots, A., Bakkerm J. and Smant, G. (2000). Degradation of plant cell walls by a nematode. Nature 406,36-37.

Petersen, T.N., Brunak, S., Von Heijne, G. and Nielsen, H. (2011). SignalP 4.0: discriminating signal peptides from transmembrane regions. Nature Methods 8(10), 785- 786.

Phillips, W.S., Howe, D.K., Brown, A.M.V., Eves-van den Akker, S., Dettwyler, L., Denver, D.R. and Zasada, I.A. (2017). The Draft Genome of Globodera ellingtonae. Journal of Nematology 49(2), 127–128.

Postma, W.J., Slootweg, E.J., Rehman, S., Finkers-Tomczak, A., Tytgat, T.O.G., van Gelderen, K., Lozano-Torres, J.L., Roosien, J., Pomp, R., van Schaik, C., Bakker, J., Goverse, A. and Smant, G. (2012). The effector SPRYSEC-19 of Globodera rostochiensis suppresses CC-NB-LRR mediated disease resistance in plants. Plant Physiology 160(2), 944-954.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Powell, S., Szklarczyk, D., Trachana, K., Roth, A., Kuhn, M., Muller, J., Arnold, R., Rattei, T., Letunic, I., Doerks, T. and Jensen, L.J. (2012). eggNOG v3. 0: orthologous groups covering 1133 organisms at 41 different taxonomic ranges. Nucleic Acids Research, 40(D1), D284-D289.

Schulze-Lefert P. and Panstruga R. (2011). A molecular evolutionary concept connecting nonhost resistance, pathogen host range, and pathogen speciation. Trends in Plant Sciences 16(3), 117–125.

Qin, L., Kudla, U., Roze, E., Goverse, A., Popeijus, H., Nieuwland, J., Overmars, H., Jones, J.T., Schots, A., Smant, G., Bakker, J. and Helder, J. (2004). A nematode expansin acting on plants. Nature 427, 30.

Raffaele, S., Farrer, R.A., Cano, L.M., Studholme, D.J., MacLean, D., Thines, M., Jiang, R.H., Zody, M.C., Kunjeti, S.G., Donofrio, N.M., Meyers, B.C., Nusbaum, C. and Kamoun, S. (2010). Genome evolution following host jumps in the Irish potato famine pathogen lineage. Science 330(6010), 1540–1543.

Replogle, A., Wang, J., Bleckmann, A., Hussey, R.S., Baum, T.J., Shinichiro, S., Davis, E.L., Wang, X., Simon, R. and Mitchum, M.G. (2011). Nematode CLE signaling in Arabidopsis requires CLAVATA2 and CORYNE. The Plant Journal 65(3), 430-440.

Replogle, A., Wang, J., Paolillo, V., Smeda, J., Kinoshita, A., Durbak, A., Tax, F.E., Wang, X., Sawa, S. & Mitchum, M.G. (2013). Synergistic interaction of CLAVATA1, CLAVATA2, and RECEPTOR-LIKE PROTEIN KINASE 2 in cyst nematode parasitism of Arabidopsis. Molecular Plant-Microbe Interactions 26(1), 87-96.

Sacco, M.A., Koropacka, K., Grenier, E., Jaubert, M.J., Blanchard, A., Goverse, A., Smant, G. and Moffett, P. (2009). The cyst nematode SPRYSEC protein RBP-1 elicits Gpa2- and RanGAP2-dependent plant cell death. PLoS Pathogens 5(8), e1000564.

Smant, G., Stokkermans, J.P., Yan, Y., de Boer, J.M., Baum, T.J., Wang, X., Hussey, R.S., Gommers, F.J., Henrissat, B., Davis, E.L., Helder, J., Schots, A. and Bakker, J. (1998). Endogenous cellulases in animals: isolation of beta-1, 4-endoglucanase genes from two species of plant-parasitic cyst nematodes. Proceedings of the National Academy of Sciences USA 95(9),4906-4911.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Simão, F.A., Waterhouse, R.M., Ioannidis, P., Kriventseva, E.V. and Zdobnov, E.M. (2015). BUSCO: assessing genome assembly and annotation completeness with single-copy orthologs. Bioinformatics 31(19), 3210-3212.

Sobczak, M. and Golinowski, W. (2011). Cyst nematodes and syncytia. In: Jones, J.T., Gheysen, G. and Fenoll, C. (eds.). Genomics and Molecular Genetics of Plant- Nematode Interactions. Springer Academic Publishers, Dordrecht, The Netherlands, pp. 61-82.

Stam, R., Mantelin, S., McLellan, H. and Thilliez, G. (2014). The role of effectors in nonhost resistance to filamentous plant pathogens. Frontiers in Plant Science 5, 582.

Smith-Unna, R., Boursnell, C., Patro, R., Hibberd, J.M. and Kelly, S. (2016). TransRate: reference-free quality assessment of de novo transcriptome assemblies. Genome Research 26(8), 1134-1144.

Trifinopoulos, J., Nguyen, L.-T., von Haeseler, A. and Minh, B.Q. (2016). W-IQ-TREE: a fast online phylogenetic tool for maximum likelihood analysis. Nucleic Acids Research, 44 (Web Server issue), W232–W235.

Vanholme, B., Haegeman, A., Jacob, J., Cannooot, B. and Gheysen, G. (2009). Arabinogalactan endo- 1,4- beta- galactosidase: a putative plant cell wall-degrading enzyme of plant parasitic nematodes. Nematology 11, 739-747.

Vogel, J. (2008). Unique aspects of the grass cell wall. Current Opinion in Plant Biology 11, 301-307.

Wang, J., Joshi, S., Korkin, D. and Mitchum, M.G. (2010). Variable domain I of nematode CLEs directs post-translational targeting of CLE peptides to the extracellular space. Plant Signaling and Behavior 5(12), 1633-1635.

Chapter 3: Adaptation to a monocot host in the plant-parasitic nematode H. sacchari

Wang, J., Lee, C., Replogle, A., Joshi, S., Korkin, D., Hussey, R., Baum, T.J., Davis, E.L., Wang, X. And Mitchum, M.G. (2010a). Dual roles for the variable domain in protein trafficking and host-specific recognition of Heterodera glycines CLE effector proteins. New Phytologist 187(4), 1003–1017.

Yang, Y. and Smith, S.A. (2013). Optimizing de novo assembly of short-read RNA-seq data for phylogenomics. BMC Genomics 14, 328.

Zhang, Y., Yang, S., Song, Y., and Wang, J. (2014). Genome-wide characterization, expression and functional analysis of CLV3/ESR gene family in tomato. BMC genomics 15(1), 827.

Zheng, M., Long, H., Zhao, Y., Lin, L., Xu, D., Zhang, H., et al. (2015) RNA-Seq based identification of candidate parasitism genes of cereal cyst nematode (Heterodera avenae) during incompatible infection to Aegilops variabilis. PLoS ONE 10(10), e0141095.

Chapter 4: General conclusion

During the course of my research work, I tried to answer following questions.

Why study rice?

Rice is a staple food for more than half of the world’s population. It is cultivated over 163 million ha area extending across 100 countries representing 30% of the world’s total cereal production (Shrestha et al., 2007; Laborte et al., 2017). If the current trend of population growth continues, an additional 116 million tons of rice will be needed to feed the growing population by 2035; meaning 8-10 million tons of additional crop per year over the next decade (Seck et al., 2012). Rice production is threatened by increasing water scarcity due to climate change. In Asia; which produces 90% of the world’s rice, it is predicted that 22 million ha rice area may face water shortages by 2025 (Tuong and Bouman 2003). With reducing water availability, bringing more land under rice cultivation is challenging; instead farmers are forced to adopt aerobic rice cultivation to tackle water crisis. Increasing rice productivity as well as reducing the losses caused by various stresses (biotic and abiotic) is crucial to achieve the goal of food security.

Why Heterodera sacchari?

Rice grown in tropical and subtropical regions of the world, is attacked by a wide range of plant pathogens in general including many PPNs. More than 150 species of PPNs infect rice and can cause annual yield losses of up to 25% (Bridge et al., 2005; Kyndt et al., 2014). Four species of cyst nematodes are reported to attack rice, of these Heterodera sacchari has the most widespread occurrence and has been reported from various countries from Asia, Africa and the Caribbean islands. H. sacchari, commonly known as the sugarcane cyst nematode is considered as an important parasite and limiting factor in sugarcane and rice production. In West Africa H. sacchari is one of the most important parasites in upland rice cultivation and can cause yield reductions of up to 50 % (Coyne and Plowright, 2000; Annonymous, 2014). A field study shows that with reduced water availability in sandy soils, H. sacchari damage in susceptible varieties is significantly increased and that the presence of nematode aggravates the effects of drought and drought-related losses (Audebert et al., 2000).

In spite of the economic importance of H. sacchari, very little is known about the details of its life cycle. At the start of this project, nothing was known about the molecular basis of the interactions between the nematode and its host. We therefore planned a series experiments in order to better understand the life cycle of H. sacchari under in vitro conditions and to better

Chapter 4: General conclusion understand the molecular basis of parasitism by this nematode. In addition to providing basic information on the nematode, by comparing our data with those generated from other cyst nematodes that parasitize dicotyledonous plants, we aimed to develop a better understanding of how host-specific pathogens become adapted to different plants.

In our initial experiments, we examined whether commonly used rice cultivars showed any differences in susceptibility to H. sacchari. These experiments showed that, all the rice cultivars tested were highly susceptible, with the highest number of cysts were recorded on Nipponbare. As Nipponbare is a commonly used model rice cultivar, we decided to use this as a susceptible host for all future experiments.

Some PPNs infect another model plant, Arabidopsis thaliana which is commonly used in plant biology, genetical and host-pathogen interaction studies. Sijmons et al. (1991), established culture conditions to study infection and development of various cyst nematodes (H. schachtii, H. trifolii and H. cajani), root-knot nematodes (M. incognita and M. arenaria) and the lesion nematode Pratylenchus penetrans under monoxenic condition using Knop medium (Sijmons et al., 1991). However, we found that Arabidopsis is not a host for H. sacchari, with strong necrosis observed at the point of invasion, ruling out the use of this plant as a model host.

To study the H. sacchari life cycle under monoxenic conditions and to avoid any contamination during the experiment, we sterilised rice seeds, growth medium and nematodes. Firstly, we assessed several different methods for rice seed sterilization. Several chemicals including ethanol, sodium/calcium hypochlorite, mercury chloride, silver nitrate and bromine water have previously been used as seed disinfectants (Oyebanji et al., 2009). The concentration and time of exposure to the selected chemicals varies depending upon nature of seed (with or without seed coat) and amount of initial contaminating inoculum the seeds carry. The method which is commonly used for Arabidopsis seed sterilization (0.6 % sodium hypochlorite followed by 70% ethanol for 5 minutes each) or the method previously described for rice seed sterilization by Kumari et al. (2016) (overnight seed soaking in distilled water with shaking followed by treatment with 70% ethanol for 30 seconds) was not enough to effectively sterilize seeds under the conditions tested here. Oyebanji et al. (2009), concluded that treating rice seeds with 90% ethanol for 3 minutes followed by 3.5 % sodium hypochlorite for 30 minutes is the most effective method for reducing contamination in rice for up to 9 days. As our experiments were planned to continue for 2 months or more, we modified this method to suit our requirements. After several trials, we were able to optimise

Chapter 4: General conclusion the seed sterilization method. The method, consisting of overnight soaking (16-18 hrs) of rice seeds in distilled water on a shaker and treatment with 70% ethanol followed by 6% sodium hypochlorite each for 15 minutes was sufficient to sterilise the majority (90%) of seeds without loss of seed viability.

The media also plays a critical role during in vitro studies of plant-nematode interactions and specific media conditions are necessary for obtaining a successful infection. For example, the sugar beet cyst nematode H. schachtii infects Arabidopsis well when the host plants are in Knop media and can be cultured and multiplied under in vitro conditions on white mustard (Sinapis alba L. cv. Albatros) plants grown aseptically on this media (Habash et al., 2017). By contrast the root-knot nematode M. incognita shows optimal infection of Arabidopsis when grown on MS-Gelrite medium as compared to Knop medium (Mendy et al., 2017).

During the in vitro experiments, we found that rice grows well on commonly used media including Knop and MS media but that J2s of H. sacchari were unable to infect rice roots growing on these media efficiently. These results suggested the need for another media for the proposed study.

Pluronic gel has been used as an alternative culture medium to agar for plant seedlings, that are subsequently infected with bacteria, fungi and nematode species (Ko and Van Gundy, 1988). Since then very few studies have been done with nematode using pluronic gel as culture medium. Pluronic gel was used to study the behavior and attraction of M. incognita towards different host plants (Wang et al., 2009); and to understand host recognition, invasion, development and reproduction behavior of two root-knot nematodes (M. incognita and M. graminicola) on rice and tomato plants (Dutta et al., 2011). In another study, pluronic gel was used as a medium to screen rice cultivars and to study infection and development of M. graminicola on resistant and susceptible rice cultivars (Kumari et al., 2016). These data suggested that pluronic gel supported growth of rice roots in a way that may allow efficient infection by H. sacchari.

We found that rice seedlings, germinated on filter paper and transplanted into 23% pluronic gel, supported high nematode infection levels and that H. sacchari completed its life cycle under these conditions in 7-9 weeks’ time. We observed and documented the complete life cycle of H. sacchari in the pluronic gel. Interestingly, unlike other cyst nematodes which reproduce by amphimixis, H. sacchari reproduces by mitotic parthenogenesis. We also found

Chapter 4: General conclusion that most of the juveniles developed into females and rarely observed males. This is in agreement with previous studies on this nematode.

The use of pluronic gel has several advantages over media plates made with agar. Unlike agar plates, the high transparency of pluronic gel makes it easy to monitor the nematode movement and host responses to infection (Wang et al., 2009; Sasaki-Crawley et al., 2012). It was previously reported that nematodes can perceive the gradients of attractants inside the pluronic gel better than on the surface of media and that this provides a more natural system for studying behavior. For example, nematodes on pluronic gel were attracted more to host roots than those of a non-host or less preferred host (Wang et al., 2009). Similarly, an attraction assay using tomato, rice and mustard roots confirms that M. incognita and M. graminicola are more attracted to roots of tomato and rice respectively than those of mustard roots (Dutta et al., 2011).

On agar plates, the roots on the surface of agar are more prone to nematode attack. By contrast in the pluronic gel system, all the host roots are equally exposed to nematode invasion mimicking soil conditions and making the assay three dimensional (Robinson, 1994; Sasaki-Crawly et al., 2012). Pluronic gel liquefies as temperature drops below a certain level dependent on the concentration of gel used. A gel of 20-25 % is in a liquid state when the temperature falls below 10°C (Gardener and Jones, 1984). This property can be exploited for synchronizing nematode infection. As pluronic gel is a non-nutrient media, it is relatively less prone to contamination with microbes and can be used without sterilization for short-term experiments. For experiments lasting more than 3 days, it can be used after sterilization by autoclaving (Ko and Van Gundy, 1988). We also found that experiments lasting more than 5 days start to get contamination if the pluronic gel was used without sterilization.

The present system can be used to study the host-pathogen interaction in detail or to collect infected root or nematode samples at different time points for various analysis. While studying the in vitro life cycle of H. sacchari on pluronic gel, we also collected 2 different stages in the life cycle of H. sacchari (2nd stage pre-parasitic juvenile and 15 days post- infection (dpi) females) for our subsequent experiments, transcriptome analysis of H. sacchari to identify important effectors.

Transcriptome Analysis

The interactions of PPNs with their host are mediated by effectors. Effectors can be defined as secreted proteins and small molecules that change host cell structure and function

Chapter 4: General conclusion

(Hogenhout et al., 2009). Effectors play a central role in parasitism and have also been referred to as parasitism genes (Davis et al., 2008). Nematode effectors have been shown to promote penetration of the root tissues, suppress the host defence and in case of sedentary parasites modify the selected root cells into the feeding structure (Curtis, 2007; Gheysen and Mitchum, 2011; Smant and Jones, 2011). The principal source of nematode effectors is the esophageal gland cells. The effectors from the gland cells are introduced into the host plant through the stylet.

Several approaches are currently used to identify effectors, and these are built on resources generated in genomics, transcriptomics and proteomics studies (Haegeman et al., 2012). The availability of genome and transcriptome data from many cyst nematode species has accelerated the process of effector identification (e.g. Gao et al., 2003; Cotton et al., 2014; Kumar et al., 2014; Zheng et al., 2015; Eves-van den Akker et al., 2016). Transcriptome analysis using 2 or more nematode life stages (preferably pre-parasitic and parasitic) or specific gland tissues is one of the best approaches to identify nematode effectors (Maier et al., 2013). Various bioinformatic tools can be used to predict the putative secretory proteins that have a signal peptide and that lack transmembrane domains. The subset of these secreted proteins that are upregulated in parasitic stages indicating their role in plant parasitism are then considered as candidate effectors that can be further studied (Thorpe et al., 2014). Expression of these proteins in the gland cells needs then to be confirmed using in situ hybridization in order to confirm that the protein is an effector.

The PPNs of highest economic importance are the sedentary endoparasites. Compared to root-knot nematodes which are polyphagous, cyst nematodes tend to have restricted host ranges. For example, potato cyst nematodes are restricted to infection of a small number of solanaceous plants while H. sacchari is only able to infect monocotyledonous plants. Although very little is known about the molecular determinants of host range in plant parasitic nematodes, effectors have been shown to have a central role in this process in other pathogens (Stam et al., 2014). A view has emerged that failure of parasitism on non-hosts that are closely related to the main host of a pathogen is most likely due to recognition of effectors by a resistance gene while a failure to infect a more distantly related species is most likely due to the inability of effectors to suppress PAMP triggered immunity (Schulze-Lefert and Panstruga, 2011). For example, comparisons of the genomes of the Irish potato famine pathogen Phytophthora infestans and a closely related species Phytophthora mirabilis, which infects Mirabilis jalapa revealed that 82 of 345 genes which showed signs of positive

Chapter 4: General conclusion selection could encode effector sequences (Raffaele et al., 2010). Subsequent work on orthologous effectors that encode protease inhibitor from the two species showed that the protease inhibitor effectors from each of these species interact specifically with protease targets from their respective host plants (Dong et al., 2014). Host-pathogen co-evolution is therefore reflected in adaptations of effectors for function in the host.

In order to gain more insights into host range determination, we choose three cyst nematode species that have different host ranges. The potato cyst nematode, G. pallida can infect and multiply on solanaceous plants; the sugarbeet cyst nematode H. schachtii has a wide host range and can multiply on plants from amaranthaceae, brassicaceae family (Turner and Subbotin, 2013). By contrast, sugarcane and rice are the only crops reported as hosts for H. sacchari. G. pallida and H. schachtii are parasite of dicots while H. sacchari is monocot parasite. Comparing the genomes or transcriptomes of these related nematode species may shed light on the genes conserved (termed core effectors) across the cyst nematodes along with genes which are more diverged or evolved to suite unique biology of each of these species. This comparison may also lead to identification of new effector candidates; as well as expansion or reduction or modification of gene families governing the host range of nematodes.

Prior to the start of this study nothing was known at the molecular level about the interaction between rice and H. sacchari. To address this, we generated transcriptome data from two different life stages (pre-parasitic J2s and 15 days post-infection) by using Illumina MiSeq next-generation sequencing generating a total of 17,086,132 paired-end 2x 250 bp reads.

After filtering with transrate and removing contamination, sequences were pooled and assembled into a single reference transcriptome consisting of 44,230 transcripts. Comparisons with published cyst nematode genome sequences showed that 34,203 (77.3%) and 30,482 (68.9%) of the H. sacchari transcripts matched sequences in G. rostochiensis and G. pallida respectively (e value 1e-10). We found that 185 out of 295 G. rostochiensis effectors identified had similar sequence in the H. sacchari transcriptome (e value <1e-10). When effector-like sequences and non-effector sequences of H. sacchari were compared with their most similar homologues in G. rostochiensis genomes, we observed that on an average, the putative effectors of H. sacchari are more different to their closest effector homologue in G. rostochiensis than non-effectors to their corresponding closest homologue. Taken together, these data show that effectors have accumulated more mutations that than non-effectors since

Chapter 4: General conclusion these species diverged. We hypothesise that this reflects the need for co-evolution with their respective hosts.

Based on comparison with other plant parasitic nematodes we choose three different groups containing putative effector proteins of H. sacchari.

The 1st group contains proteins which are not usually found in animals and are believed to have been acquired from various prokaryotes (mainly bacteria) through horizontal gene transfer events (Danchin et al., 2010). Many of these proteins play an important role in nematode penetration and migration inside the host roots. From the H. sacchari dataset we identified sequences similar to glycosyl hydrolase family 5 (GHF5) cellulase which acts on cellulose; pectate lyase capable of degrading pectate sugar polymers; expansins and cellulose binding proteins which soften the cell wall and enhances the activity of other hydrolytic enzymes.

Analysis of the expression profile of one of the glycosyl hydrolase family 5 (GHF5) cellulases identified from the H. sacchari dataset, showed that it is highly upregulated in pre- parasitic juveniles compared to other stages (eggs, 15 and 25 dpi). In situ hybridization of pre-parasitic juveniles using Digoxigenin (DIG)-labelled probes shows that its expression is restricted to subventral esophageal gland cells which is consistent with previous observations of GHF5 cellulases from H. glycines (Smant et al., 1998) and G. rostochiensis (Rehman et al., 2009a) and other nematodes. It is known that the subventral gland cells are highly active during the early stage of infection and secretes enzymes used for penetration and migration (Davis et al., 2008); most of the genes acting on the cell wall are expressed in the subventral gland cells (Haegeman et al., 2012).

In addition to these genes, we also found GH32 invertases which metabolise sucrose and two sequences similar to chorismate mutase. Each of these sequences has been previously identified in other PPNs. Like cellulase, we also did the expression analysis and in situ hybridization for one of the transcripts similar to chorismite mutase. Our results show that chorismate mutase is expressed in all the life cycle stages tested and in situ hybridization confirms its presence in esophageal glands. These results are in line with other studies where Vanholme et al. (2009), investigated chorismite mutase from H. schachtii and found that chorismate mutase gens is expressed in pre-parasitic and parasitic stages of nematode and localized in both the gland cells while subventral gland cells specific expression was found in G. pallida (Jones et al., 2003). The function of this effector has been analysed in other PPNs

Chapter 4: General conclusion and it may reduce the availability of chorismate for conversion into plant defence compounds (Doyle and Lambert, 2003; Jones et al., 2003; Vanholme et al., 2009).

We also found that the H. sacchari and H. avenae transcriptomes do not contain GH53 (endo-beta-1,4-galactanase) enzymes. The GH53 enzyme is present in all other cyst nematodes analysed so far which parasitize dicot plants. One possible explanation for the absence of GH53 in the monocot parasites (H. sacchari and H. avenae) is the composition of the cell wall of monocot roots. The primary cell wall of monocot roots contains relatively fewer proportions of pectic polysaccharides which is the substrate of GH53 enzymes (Vogel, 2008). The absence of GH53 enzyme may show nematode adaptation to parasitism of monocot plants.

Another important family of candidate effector are the SPRYSECs, which stands for secreted SPRY domain-containing proteins. The SPRY domain was first identified from Dictyostelium discoideum in the dual specificity kinase spore lysis SP1a protein and in three mammalian Ca2+-release channel RYanodine receptors (Ponting et al., 1997). The SPRYSEC effector family was initially identified from cyst nematodes (Qin et al., 2000; Blanchard et al., 2005). Some SPRYSEC effectors (but not all) from G. pallida and G. rostochiensis are involved in host defence suppression (Diaz-Granados et al., 2016; Mei et al., 2015). The genome of G. pallida contains 295 SPRY domain containing sequences of which 30 have a predicted signal peptide while G. rostochiensis has 71 SPRY domain containing sequences of which 17 have a signal peptide (Cotton et al., 2014; Eves-van den Akker et al., 2016). This data suggests that even though the SPRYSEC family is highly expanded in cyst nematodes, only a minority of the SPRY domain proteins have a predicted signal peptide for secretion thus allowing their deployment as an effector (Mei et al., 2015). Most of the SPRYSEC effector identified to date are from the potato cyst nematodes and found to be expressed in dorsal esophageal gland cells (Blanchard et al., 2005; Jones et al., 2009; Rehman et al., 2009).

The Gp-RBP1, the secreted protein from G. pallida has a SPRY domain which closely resembles the Ran-Binding Protein in the Microtubule-organizing center (RanBPM) (Rehman et al., 2009). Several variants of Gp-Rbp-1 from G. pallida have been identified and showed a high degree of polymorphism in both virulent and avirulent population to Gpa2. The Gpa2 is a potato resistance gene that encodes a CC-NB-LRR protein. The potato plants expressing Gpa2 are able to recognize Gp-RBP1 from all avirulent G. pallida populations and elicit defense response in the form of cell death. This recognition of Gp-RBP1 by Gpa2 is enabled by a single amino acid polymorphism at position 187 in the Gp-RBP-1 SPRY domain.

Chapter 4: General conclusion

Substitution of proline to serine abolishes recognition. While serine to proline substitution gains the recognition and confers resistance against G. pallida (Van der Vossen et al., 2000; Sacco et al., 2009). In another study, Rehman et al. (2009), used yeast-two-hybrid (Y2H) screening to confirm the interaction of SPRYSEC-19 of G. rostochiensis with a CC-NB-LRR protein in an SW5 resistant gene cluster and postulates that this interaction may probably downregulate immune activity in susceptible tomato plants (Rehman et al., 2009).

The H. sacchari dataset contained 80 transcripts encoding putative SPRY domain proteins, 6 of which have a predicted N terminal signal peptide allowing their secretion. As in the case of PCN, SPRYSECs from H. sacchari were found to be expressed in the dorsal pharyngeal gland cell of J2s. However, two transcripts analysed by qPCR shows upregulation in the parasitic stages which is in contrast with SPRYSECs from PCNs which are upregulated in pre-parasitic and parasitic J2s (Rehman et al., 2009; Mei et al., 2015). The role of SPRYSEC effectors in H. sacchari remains to be tested.

The 3rd group of effectors which we studied are important in the formation and maintenance of nematode feeding site and which belong to CLAVATA3/EMBRYO SURROUNDING REGION-like (CLE) family. We also studied the H. sacchari orthologue of the 19C07 effector.

CLE family peptides have been reported from several monocot and dicot plants as well as plant-parasitic nematodes. In plants, CLEs regulate plant growth and development including shoot, root and floral meristem maintenance, vascular development, organ size regulation and apical dominance (Cock and McCormick, 2001; Sawa et al., 2008; Yamada and Sawa, 2013).

Several CLE genes have been cloned from cyst nematodes including H. glycines, G. rostochiensis and H. schachtii (Wang et al., 2005; Lu et al., 2009; Wang et al., 2011). The general structure of CLE proteins from nematodes has an N-terminal signal peptide followed by an internal variable domain and conserved single or multiple CLE domain(s) at the C- terminal (Cock and McCormick, 2001). Wang et al. (2011), cloned two CLE like genes (HsCLE1 & HsCLE2) from H. schachtii and found that the 12 amino acid CLE motif from HsCLE2 was identical to Arabidopsis AtCLE5 and AtCLE6 CLE genes. They also reported that overexpression of HsCLEs in Arabidopsis resulted in a wuschel-like phenotype (Wang et al., 2011), while the exogenous application of G. rostochiensis CLE peptide gives short root phenotype in potato plants (Chen et al., 2015). It was also reported that an Arabidopsis mutant can be rescued by expressing CLE gene from nematode (Wang et al., 2005; Lu et al.,

Chapter 4: General conclusion

2009). Its concluded from the various experiments that nematode CLEs can mimic the plant CLEs and alter or re-programme the host root cell for the feeding site formation (Wang et al., 2011).

The transcriptome of H. sacchari has seven transcripts containing a putative CLE domain. H. sacchari CLEs also encodes an N-terminal signal peptide. 6 out of 7 transcripts present in H. sacchari have single canonical CLE domain at C terminus while the seventh transcript has a tandemly repeated motif without clear homology to canonical CLE domains. A comparison of plant and nematode CLEs showed that H. sacchari CLEs are more similar to rice CLEs than to those from other plants or other plant-parasitic nematodes, suggesting functional adaptation to parasitism of rice. This was confirmed in functional studies in which we showed that the exogenous application of H. sacchari CLEs resulted in shoot root phenotype in rice whereas peptides from H. glycines or H. sacchari shuffled peptide had no effect.

In H. sacchari transcriptome dataset, we also found one transcript similar to the 19C07 effector of H. schachtii. The H. schachtii 19C07 effector has been found to interact with auxin transporter gene, LAX3 in Arabidopsis. This interaction increases auxin influx inside the root cell which may stimulate the syncytium development (Lee et al., 2011). Hs19C07, when cobombarded with LAX3 using BiFC vector system, interaction was found at the periphery of onion epidermal cell. Although the interaction between H. sacchari 19C07 with LAX3 gene of rice is yet to studied, the transient expression of H. sacchari 19C07 in the leaves of N. benthamiana showed its localization in the cell periphery (Fig. S1), suggesting the function may be conserved across the species.

To conclude, we found that pluronic gel is good medium to study rice-H. sacchari interaction. We have generated transcriptomic data of H. sacchari which will be a valuable resource for comparative analysis and future genomic studies. We have used these data to show that effectors from H. sacchari are adapted to allow parasitism of rice.

Chapter 4: General conclusion

Fig 4. S1: Transient expression of H. sacchari 19C07 in the leaves of N. benthamiana showed its localization in the cell periphery. a) Green fluorescence originated from H. sacchari 19C07: GFP fusion protein. b) Purple fluorescence originated from apoplastic marker:mCherry fusion protein.

4.2 References

Annonymous (2014). Invasive species compendium, CABI’s report for sugarcane cyst nematode, Heterodera sacchari. (http://download.ceris.purdue.edu/file/3047).

Audebert, A., Coyne, D.L., Dingkuhn, M. and Plowright, R.A. (2000). The influence of cyst nematodes (Heterodera sacchari) and drought on water relations and growth of upland rice in Côte d’Ivoire. Plant and soil 220(1-2), 235–242.

Blanchard, A., Esquibet, M., Fouville, D. and Grenier, E. (2005). Ranbpm homologue genes characterised in the cyst nematodes Globodera pallida and Globodera mexicana. Physiological and Molecular Plant Pathology 67(1),15-22.

Bridge, J., Plowright, R.A. and DeLiang, Peng. (2005). Nematode parasites of rice. In: Luc, M., Sikora, R.A. & Bridge, J. (edss). Plant parasitic nematodes in subtropical and tropical agriculture. CABI, pp. 87–128.

Chen, S. and Wang, X. (2015). In vitro CLE peptide bioactivity assay on plant roots. Bio- protocol 5(24), 1-6.

Cock, J.M. and McCormick, S. (2001). A large family of genes that share homology with CLAVATA3. Plant Physiology 126, 939–942.

Chapter 4: General conclusion

Globodera pallida elucidate key aspects of plant parasitism by a cyst nematode. Genome Biology 15(3), R43.

Coyne, D.L. and Plowright, R.A. (2000). Pathogenicity of cyst nematode, Heterodera sacchari, on rice in sand and clay soil. International Rice Research Notes 25 (1), 17–18.

Curtis, R.H.C. (2007). Plant parasitic nematode proteins and the host-parasite interaction. Briefings in Functional Genomics and Proteomics 6(1), 50–58.

Diaz-Granados, A., Petrescu, A.J., Goverse, A. and Smant, G. (2016). SPRYSEC Effectors: A versatile protein-binding platform to disrupt plant innate immunity. Frontiers in Plant Science 7, 1575.

Davis, E.L., Hussey, R.S., Mitchum, M.G. and Baum, T.J. (2008). Parasitism proteins in nematode-plant interactions. Current Opinion in Plant Biology 11(4), 360–366.

Dong, A., Stam, R., Cano, L.M., Song, J., Sklenar, J., Yoshida, K., Bozkurt, T.O., Oliva, R., Liu, Z., Tian, M., Win, J., Banfield, M.J., Jones, A.M.E., van der Hoorn, R.A.L. and Kamoun, S. (2014). Effector specialization in a lineage of the Irish potato famine pathogen. Science 343(6170), 552-555.

Doyle, E.A. and Lambert, K.N. (2003). Meloidogyne javanica chorismate mutase 1 alters plant cell development. Molecular Plant-Microbe Interactions 16, 123–131.

Chapter 4: General conclusion

Phillips, W., Henrissat, B., Urwin, P.E., Blaxter, M. and Jones, J.T. (2016). The genome of the yellow potato cyst nematode, Globodera rostochiensis, reveals insights into the bases of parasitism and virulence. Genome Biology 17, 124.

Gao, B., Allen, R., Maier, T., Davis, E.L., Baum, T.J. and Hussey, R.S. (2003). The parasitome of the phytonematode Heterodera glycines. Molecular Plant-Microbe Interactions 16(8), 720–726.

Gardener, S. and Jones, J.G. (1984). A new solidifying agent for culture media which liquefies on cooling. Journal of General Microbiology 130, 731–733.

Gheysen, G. and Mitchum, M.G. (2011). How nematodes manipulate plant development pathways for infection. Current Opinion in Plant Biology 14(4), 415–421.

Habash, S.S., Sobczak, M., Siddique, S., Voigt, B., Elashry, A. and Grundler, F.M.W. (2017). Identification and characterization of a putative protein disulfide isomerase (HsPDI) as an alleged effector of Heterodera schachtii. Scientific reports 7(1), 13536.

Haegeman, A., Mantelin, S., Jones, J.T. and Gheysen, G. (2012). Functional roles of effectors of plant-parasitic nematodes. Gene 492(1), 19–31.

Hogenhout, S.A., Van der Hoorn, R.A.L., Terauchi, R. and Kamoun, S. (2009). Emerging concepts in effector biology of plant-associated organisms. Molecular Plant-Microbe Interactions 22,(2) 115–122.

Jones, J.T., Furlanetto, C., Bakker, E., Banks, B., Blok, V., Chen, Q., Phillips, M. and Prior, A. (2003). Characterization of a chorismate mutase from the potato cyst nematode Globodera pallida. Molecular plant pathology 4(1), 43-50.

Ko, M.P. and Van Gundy, S.D. (1988). An alternative gelling agent for culture and studies of nematodes, bacteria, fungi, and plant tissues. Journal of Nematology 20(3):478-485.

Kumar, M., Gantasala, N.P., Roychowdhury, T., Thakur, P.K., Banakar, P., Shukla, R.N.,

Chapter 4: General conclusion

Jones, M.G.K., Rao, U. (2014). De novo transcriptome sequencing and analysis of the cereal cyst nematode, Heterodera avenae. PLOS One 9(5): e96311.

Kyndt, T., Fernandez, D. and Gheysen, G. (2014). Plant-parasitic nematode infections in rice: Molecular and cellular insights. Annual Review of Phytopathology 52, 135–153.

Laborte, A.G., Gutierrez, M.A., Balanza, J.G., Saito, K., Zwart, S.J., Boschetti, M., Murty, M.V.R., Villano, L., Aunario, J.K., Reinke, R., Koo, J., Hijmans, R.J. and Nelson, A. (2017). RiceAtlas, a spatial database of global rice calendars and production. Scientific Data 4:170074.

Mei, Y., Thorpe, P., Guzha, A., Haegeman, A., Blok, V.C., MacKenzie, K., Gheysen, G., Jones, J.T. and Mantelin, S. (2015). Only a small subset of the SPRY domain gene family in is likely to encode effectors, two of which suppress host defences induced by the potato resistance gene. Nematology 17(4), 409–424.

Mendy, B., Wang’ombe, M.W., Radakovic, Z.S., Holbein, J., Ilyas, M., Chopra, D., Holton, N., Zipfel, C., Grundler, F.M.W. and Siddique, S. (2017). Arabidopsis leucine-rich repeat receptor–like kinase NILR1 is required for induction of innate immunity to parasitic nematodes. PLOS Pathogens 13(4), e1006284.

Chapter 4: General conclusion

Oyebanji, O.B., Nweke, O., Odebunmi, O., Galadima, N.B., Idris, M.S., Nnodi, U.N., Afolabi, A.S. and Ogbadu, G.H. (2009). Simple, effective and economical explant- surface sterilization protocol for cowpea, rice and sorghum seeds. African Journal of Biotechnology 8 (20), 5395-5399.

Ponting, C., Schultz, J. and Bork, P. (1997). SPRY domains in ryanodine receptors (Ca2+- release channels). Trends in Biochemical Science 22(6), 193-194.

Qin, L., Overmars, H., Helder, J., Popeijus, H., van der Voort, J.R., Groenink, W., van Koert, P., Schots, A., Bakker, J. and Smant, G. (2000). An Efficient cDNA-AFLP-based strategy for the identification of putative pathogenicity factors from the potato cyst nematode Globodera rostochiensis. Molecular Plant Microbe Interactions 13(8), 830– 836.

Rehman, S., Postma, W., Tytgat, T., Prins, P., Qin, L., Overmars, H., Vossen, J., Spiridon, Laurentiu-N., Petrescu, Andrei-J., Goverse, A., Bakker, J. and Smant G. (2009). A secreted SPRY domain-containing protein (SPRYSEC) from the plant-parasitic nematode Globodera rostochiensis interacts with a CC-NB-LRR protein from a susceptible tomato. Molecular Plant Microbe Interactions 22(3), 330–340.

Rehman, S., Butterbach, P., Popeijus, H., Overmars, H., Davis, E.L., Jones, J.T., Goverse, A., Bakker, J. and Smant, G. (2009a). Identification and characterization of the most abundant cellulases in stylet secretions from Globodera rostochiensis. Phytopathology 99(2), 194-202.

Robinson, A.F. (1994). Movement of five nematode species through sand subjected to natural temperature gradient fluctuations. Journal of Nematology 26(1), 46-58.

Sasaki-Crawley, A., Curtis, R., Birkett, M., Papadopoulos, A., Blackshaw, R. and Pickett, J.

Chapter 4: General conclusion

(2012). The use of Pluronic F-127 to study the development of the potato cyst nematode, Globodera pallida. Nematology 14(7), 869–873.

Sawa, S., Kinoshita, A., Betsuyaku, S. and Fukuda, H. (2008). A large family of genes that share homology with CLE domain in arabidopsis and rice. Plant Signaling & Behavior 3(5), 337-339.

Schulze-Lefert P. and Panstruga R. (2011). A molecular evolutionary concept connecting nonhost resistance, pathogen host range, and pathogen speciation. Trends in Plant Science 16(3), 117–125.

Seck, P.A., Diagne, A., Mohanty, S. and Wopereis, M.C.S. (2012). Crops that feed the world: Rice. Food Security 4(1), 7–24.

Shrestha, R., Uzzo, F., Wilson, M.J. and Price, A.H. (2007). Physiological and genetic mapping study of tolerance to root-knot nematode in rice. New Phytologist 176(3), 665– 672.

Sijmons, P.C., Grundler, F.M.W., Von Mende, N., Burrows, P.R. and Wyss, U. (1991). Arabidopsis thaliana as a new model host for plant‐parasitic nematodes. The Plant Journal 1(2), 245–254.

Smant, G. and Jones, J.T. (2011). Suppression of plant defences by nematodes. In: Jones, J., Gheysen, G., and Fenoll, C., (eds.). Genomics and molecular genetics of plant-nematode interactions. pp. 273–286. Dordrecht: Springer Netherlands.

Stam, R., Mantelin, S., McLellan, H. and Thilliez, G. (2014). The role of effectors in nonhost resistance to filamentous plant pathogens. Frontiers in Plant Science 5, 582.

Thorpe, P., Mantelin, S., Cock, P.J.A., Blok V.C., Coke, M.C., Eves-van den Akker, S., Guzeeva, E., Lilley, C.J., Smant, G., Reid, A.J., Wright, K.M., Urwin, P.E. and Jones, J.T. (2014). Characterisation of the full effector complement of the potato cyst nematode

Chapter 4: General conclusion

Globodera pallida. BMC Genomics 15, 923.

Tuong T.P. and Bouman, B.A.M. (2003). Rice production in water-scarce environments. In: Kijne, J.W., Barker. R. and Molden, D. (eds.). Water productivity in agriculture: limits and opportunities for improvement. pp. 53–67. CABI,.Wallingford, UK.

Turner, S.J. and Subbotin, S.A. (2013) Cyst nematodes. In: Perry, R.N. and Moens, M. (eds.) Plant Nematology, 2nd edn. pp. 109–143. CABI, Wallingford, UK.

Vanholme, B., Kast, P., Haegeman, A., Jacob, J., Grunewald, W., Gheysen, G. (2009). Structural and functional investigation of a secreted chorismate mutase from the plant- parasitic nematode Heterodera schachtii in the context of related enzymes from diverse origins. Molecular plant pathology 10(2), 189-200.

Van Der Vossen, E.A.G., Van Der Voort, J.N.A.M., Rouppe, K., Konstantin, B., Abdel, S., Hans, B., David, C., Bakker, J., Stiekema, W.J. and Klein-Lankhorst, R.M. (2000). Homologues of a single resistance-gene cluster in potato confer resistance to distinct pathogens: A virus and a nematode. The plant journal 23(5) 567-576.

Vogel, J. (2008). Unique aspects of the grass cell wall. Current Opinion in Plant Biology 11, 301-307.

Wang, C., Lower, S. and Williamson, V.M. (2009). Application of pluronic gel to the study of root-knot nematode behaviour. Nematology 11(3), 453–464.

Wang, J., Replogle, A., Hussey, R., Baum, T., Wang, X., Davis, E.L. and Mitchum M.G. (2011). Identification of potential host plant mimics of CLAVATA3/ESR (CLE)-like peptides from the plant-parasitic nematode Heterodera schachtii. Molecular plant pathology 12(2), 177–186.

Yamada, M. and Sawa, S. (2013). The roles of peptide hormones during plant root development. Current Opinion in Plant Biology 16(1), 56–61.

Zheng, M., Long, H., Zhao, Y., Lin, L., Xu, D., Zhang, H., Liu, F., Deng, G., Pan, Z. and Yu,

100

Chapter 4: General conclusion

M. (2015). RNA-Seq based identification of candidate parasitism genes of cereal cyst nematode (Heterodera avenae) during incompatible infection to Aegilops variabilis. PLoS ONE 10: e0141095.

101

Acknowledgement

First of all, I would like to express my sincere gratitude to my supervisor Prof. Dr. Florian Grundler, Molecular Phytomedicine-INRES, University of Bonn for accepting me as a PhD student in his lab with excellent research facilities and international environment. His continuous support on both scientific and social level was endless. It was a great learning experience for almost three and half year. Thank you very much for the opportunity.

I would also like to express my greatest appreciation to my co-supervisor Prof. Dr. John Jones, James Hutton Institute, Dundee for his remarkable mentorship, generous support and guidance during my PhD. He was the one who redirected me to Germany for my studies also hosted me at JHI, Dundee for six months to do part of my research in his lab. I enjoyed great scientific discussions, encouragement and advices during my Ph.D.

I am grateful to Dr. Abdelnaser Elashry for his guidance during this work with initial data analysis and designing the experiments and for his kind support during my stay in Bonn. My sincere thanks goes to Dr. Sebastian Eves-van den Akker, group leader of plant- parasite/pathogen interaction group at University of Cambridge and Dr. Peter Thorpe, bioinformatician at JHI for their support during bioinformatic analysis of transcriptome data and immense contribution during manuscript preparation. Special thanks to Dr. Shahid Siddique, Prof. Dr. Michel Frei, Prof. Dr. Tina Kyndt, Prof. Dr. Godelieve Gheysen and Amalia Diaz-Granados for giving me opportunity to contribute in their research work.

It gives me great pleasure to acknowledge all my mpm colleagues for their support, advice and all the fun during my stay in Bonn with special mention of Dr. Samer Habash and Dr. Zoran Radakovic (Dr is not for me) for teaching me various techniques and helping me during my research. I enjoyed all the discussion and working in the late evenings and on weekends. Many thanks goes to Gisela Sichtermann for maintaining the H. sacchari culture in the green-house and to Stefan Neumann, Ute Schlee, for their excellent technical and Brigit Otte for administrative support.

I am highly indebted to Indian Council of Agricultural Research (ICAR), Ministry of Agriculture, Govt. of India and the Royal society, UK for financial support during my Ph.D and National Rice Research Institute (ICAR-NRRI), Cuttack Odisha for granting me study leave to pursue my dream. Many thanks to all my seniors, colleagues and friends from NRRI, Cuttack for their help and support with Dr. Totan Adak as my link to NRRI during last three and half years.

102

Acknowledgement

Also, I would like to thank the Prof. Dr Michel Frei and Prof. Dr Matthias Wüst for their support and taking part at my thesis defense.

Last but by no means least, I would like to thank all my family members (Pokhare and Patil) and friends for their support during my stay in Germany with special mention of my wife Devata for her endless and unconditional love and support and taking care of our princess in my absence.

Lastly, my sincere thanks for everyone for being with me.

Thank you one and all.

103