Investigations on the Early Stages of Interactions Between the Nematodes Meloidogyne Javanica and Pratylenchus Thornei and Two of Their Plant Hosts

Investigations on the early stages of interactions between the nematodes Meloidogyne javanica and Pratylenchus thornei and two of their plant hosts

SOSAMMA PAZHAVARICAL

Doctor of Philosophy

August 2009

University of Western Sydney

“O LORD, How great are your works!” Psalm 92:5

Dedicated to

My father the late P. V. Easow, who passed away in 1976 when I was still in high school; my brother the late P. Vergis, who passed away in early 2009 and my loving mother Mariamma Easow, who is, always, my main source of inspiration .

ACKNOWLEDGEMENTS

This thesis was one which was almost never written but for the grace of God. I wish to express my gratitude to the many who directly and indirectly contributed towards the completion of this thesis from the beginning, eight years ago.

First and foremost I would like to thank the University of Western Sydney for the Post Graduate Award Scholarship, and the Australian National University for the additional funding to enable me to realise my research dreams at the UWS Hawkesbury and ANU Canberra.

I express my deep appreciation for the understanding and support of my supervisory panel, especially my Principal Supervisor Associate Professor Robert Spooner-Hart, without whose infinite patience, dedication, lateral thinking and valuable time which he spent editing, this thesis would never have been realised.

My heartfelt thank you goes to Associate Professor Tan Nair for his valuable guidance during the initial stages of my candidature, and his advice in all academic matters up until recent retirement.

I would like to thank Professor Dr. Geoff Wasteneys for his kindness and support in providing all laboratory facilities for my cytoskeletal research at the Research School of Biological Sciences, Australian National University, Canberra. I consider it a privilege to have worked with Dr Wasteneys on my project.

I want to thank Dr. David Collings for sharing with me his technical expertise in confocal microscopy procedures, his knowledge and enthusiasm, patience and generosity through the years, and for his valuable time editing the cytoskeleton chapter. Dr Collings was the best mentor I could have ever wished to work with.

My gratitude goes to Dr Zhaohui Wang, Murdoch University Perth, WA for graciously teaching me the aseptic culturing technique of tomato and Meloidogyne javanica and also supplying me with aseptic Meloidogyne javanica inoculum; Dr Sharyn Taylor, SARDI, SA for supplying the Pratylenchus thornei inoculum; Dr Jennifer Cobon, DPI, QLD for providing additional Meloidogyne javanica inoculum, and tomato seeds for my experiment and Daigo Takemoto and Dr David Jones at RSBS, ANU for kindly supplying GFP-hTalin transformed Arabidopsis thaliana seeds for the GFP experiment.

Thank you to all other academic, technical and administrative staff from UWS, Hawkesbury and RSBS, ANU especially Oleg Nicetic, Liz Kabanoff, Rosalie Laing, Gillian Wilkins, Christina Harvey and Dr. Paul Holford at UWS and everyone from the Wasteneys’ and David Jones’ laboratories at RSBS, ANU.

I would like to thank Dr Shelley Burgin for her understanding and compassion and providing me with many extensions of candidature so that I could finish the thesis.

I would like to express my deep love and unending gratitude to my mother, Mariamma Easow, a retired science teacher, for sparking my interest in science from a very early age, my late father, P. V. Easow, a Finance Accountant, and my late brother, P. Vergis, who always emphasised the importance of a good education in my life.

I would like to thank my sister, Dr Marykutty Samuel, for her unwavering support through all the trials and tribulations of the darkest times in my life which I faced during the past few years, for encouraging me to persist in my studies when I was ready to give up on several occasions, for several reasons.

I would like to thank my sons Arun and Ashish, who have supported me and always encouraged me to finish my studies in spite of the amount of time I spent away from them for the sake of research and suffering my ill temper at times because of the stress.

I also want to convey my deepest gratitude to my sister-in-law, Lysamma Vergis, and my nieces Sherly and Sherin in India, for their love and hospitality during my visits.

I would like to express my sincere appreciation and also a very belated thank you to Shah who greatly assisted me through my Masters Degree in India and who I, regretfully, failed to acknowledge at that time.

I want to express my warm gratitude to Dr. Ahmed Regina and her family for their hospitality, friendship and kindness especially during the time of my stay in Canberra.

Thanks also to my many relatives, friends and acquaintances, too numerous to mention by name, who have knowingly or unknowingly given me strength to face difficult situations in life and an ability to see the bright side of all things

And I thank God for RG, who has shown me positivity, hope and a new perspective.

STATEMENT OF AUTHENTICATION

The work presented in this thesis is, to the best of my knowledge and belief, original except as acknowledged in the text. I hereby declare that I have not submitted this material, either in full or in part, for a degree at this or any other institution.

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(Signature)

TABLE OF CONTENTS

Page No i Table of Contents

List of Tables vii

List of Figures viii

Abbreviations xii

Summary xiv

Chapter 1 General introduction

1.1 Choosing nematodes for the study 1

1.1.1 Economic importance of Meloidogyne spp. 1 and Pratylenchus spp. in crop production

1.1.2 Control of plant parasitic nematodes 20

1.2 Choosing model host plants for this study 24

1.3 Aims and Objectives 26

1.3.1 Aim of this study 26

1.3.2 Rationale behind the aims and objectives 26 of this study

Chapter 2 Changes in the cytoskeleton in root cells of Arabidopsis thaliana due to infection by Meloidogyne javanica and Pratylenchus thornei

2.1 Introduction: The cytoskeleton in animals and 28 plants

2.1.1 Functions of the plant cytoskeleton 32

2.1.2 Dynamics and organisation of the 34 microtubules within plant cells

2.1.3 Interaction between actin and 35 microtubules in plant cells

2.1.4 Changes in the plant cytoskeleton due to 36 external factors

2.1.5 Immunofluorescence and the study of 39 plant cytoskeleton

2.1.6 GFP in the study of plant cytoskeleton 40

2.1.7 Nematodes and the plant cytoskeleton 42

2.2 Materials and Methods 44

2.2.1 Culture of Meloidogyne javanica 44 inoculum

2.2.2 Pratylenchus thornei inoculum 45

2.2.3 Culture of Arabidopsis thaliana ecotype 46 Columbia as host

2.2.4 Immunolabelling for confocal 46 microscopy of whole roots

2.2.5 Embedding and sectioning of whole roots 47 in BMM

2.2.6 Immunolabelling BMM sections for 48

confocal microscopy

2.2.7 Visualisation of the actin cytoskeleton in 48 living root cells of Arabidopsis thaliana using GFP-hTalin fusion protein

2.3 Results

2.3.1 Uninfected Arabidopsis thaliana roots 49

2.3.2 Arabidopsis thaliana infected by 62 Meloidogyne javanica

2.3.3 Arabidopsis thaliana infected by 89 Pratylenchus thornei

2.4 Discussion 92

2.5 Conclusion 98

Chapter 3 Effect of the nematodes Meloidogyne javanica and Pratylenchus thornei on growth and yield of tomato, Solanum lycopersicum

3.1 Introduction 99

3.2 Materials and Methods 100

3.2.1 Statistical analysis 101

3.3 Results 102

3.3.1 Observations on the growth stages of 102 tomato, Solanum lycopersicum, in the greenhouse

3.3.2 Meloidogyne javanica 102

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3.3.3 Pratylenchus thornei 108

3.3.4 Comparison of growth and yield of 114 tomato, Solanum lycopersicum, inoculated with Meloidogyne javanica and Pratylenchus thornei

3.4 Discussion 116

3.5 Conclusions 122

Chapter 4 Movement of Meloidogyne javanica and Pratylenchus thornei through sandy, sandy clay loam and clay soils to the roots of tomato, Solanum lycopersicum

4.1 Introduction

4.1.1 Soil types and soil texture 124

4.1.2 Organic matter in soils 125

4.1.3 Pore space and aeration in soils 126

4.1.4 Nematode morphology 126

4.1.5 Relationship between soil and movement 127 of nematodes

4.2 Materials and Methods 129

4.2.1 Statistical analysis 131

4.3 Results 131

4.3.1 Movement of Meloidogyne javanica to 131 the roots of tomato, Solanum lycopersicum, through sandy, sandy clay loam and clay soils

4.3.2 Movement of Pratylenchus thornei to the 133 roots of tomato, Solanum lycopersicum, through sandy, sandy clay loam and clay soils

4.4 Discussion 135

4.5 Conclusions 139

Chapter 5 Temporal study of the early infection of tomato, Solanum lycopersicum, by Meloidogyne javanica and Pratylenchus thornei

5.1 Introduction 141

5.2 Materials and methods 143

5.3 Results 144

5.3.1 Early infection of tomato, Solanum 144 lycopersicum root by Meloidogyne javanica

5.3.2 Early infection of tomato, Solanum 157 lycopersicum, root by Pratylenchus thornei

5.4 Discussion 164

5.5 Conclusions 167

Chapter 6 General discussion 6.1 Major findings / outcomes 168

6.1.1 Comparison between the effects of 171 Meloidogyne javanica and Pratylenchus

thornei on Arabidopsis thaliana and Solanum lycopersicum

6.1.2 Understanding the effects of nematode 172 and plant interactions

6.1.3 Soil characteristics and plant growth 176 parameters

6.1.4 Temporal study of nematode-plant 178 parasite-host relationships

6.2 Usefulness / implications of the work to others 179

6.3 Integrated management of nematode pests 180

6.4 Impact of climate change on plant-parasitic 181 nematodes

6.5 Future research work 182

6.6 Final conclusion 183

Chapter 7 References 184

Appendix 1 Results of analysis of soils used in pot 221 culture trial

LIST OF TABLES

Page Number Table 3.1 Summary of the statistical analysis for the mean shoot 103 length, dry weight of shoot and the mean number of green leaves at harvest of tomato inoculated with Meloidogyne javanica

Table 3.2 Summary of the statistical analysis for the mean dry weight 105 of root and mean number of flowers and buds at harvest of tomato plants inoculated with Meloidogyne javanica

Table 3.3 Summary of the statistical analysis for the mean diameter of 107 largest mature fruit, fresh weight of fruit at harvest, dry weight of fruit at harvest and moisture content of fruit at harvest of tomato inoculated with Meloidogyne javanica

Table 3.4 Summary of the statistical analysis for the mean shoot 109 length, dry weight of shoot and mean number of green leaves of tomato inoculated with Pratylenchus thornei

Table 3.5 Summary of the statistical analysis for the mean dry weight 111 of root and number of flowers and buds at harvest of tomato inoculated with Pratylenchus thornei

Table 3.6 Summary of the statistical analysis for the mean diameter of 113 largest mature fruit, fresh weight of fruit at harvest, dry weight of fruit at harvest and moisture content of fruit at harvest of tomato inoculated with Pratylenchus thornei

Table 3.7 Percentage reduction of the dry weights of shoot, root and 115 fruit, number of combined flowers and flower buds and fresh weight of fruit, from control

Table 3.8 Developmental stages and their durations in tomato , 117 Solanum lycopersicum

Table 4.1 Physical and chemical properties of sand, sandy clay loam 130 and clay soils used in the nematode movement experiment

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LIST OF FIGURES

Page Number Figure 2.01 The structure of actin microfilaments 29

Figure 2.02 The structure of microtubules 30

Figure 2.03A&B An Arabidopsis thaliana root triple labelled for 51&52 actin, microtubules and DNA shows the different stages of cell division in an uninfected root meristem.

Figure 2.04A&B An uninfected Arabidopsis thaliana root shows 53&54 cytoskeletal organisation in the post-mitotic root tip.

Figure 2.05 The organisation of actin and microtubules in 56 differentiating cells of an uninfected Arabidopsis thaliana root.

Figure 2.06A&B Immunolabelling of a BMM section of an 57&58 uninfected Arabidopsis thaliana root.

Figure 2.07 A GFP-hTalin-transformed Arabidopsis thaliana 60 seedling showing actin in the developing vasculature of the root tip.

Figure 2.08 GFP-hTalin labelling revealed an extensive actin 61 cytoskeleton in the epidermis of the root differentiation zone.

Figure 2.09 Immunolabelling of a Meloidogyne-infected 63 Arabidopsis root demonstrated cytoskeletal disruption during nematode entry.

Figure 2.10 Immunolabelling of a Meloidogyne-infected 64 Arabidopsis root demonstrated cytoskeletal disruption adjacent to the root tip during nematode entry.

Figure 2.11 An Arabidopsis thaliana root tip infected with 65 Meloidogyne javanica showed increased cytoskeletal labelling and disruption.

Figure 2. 12 Cytoskeletal changes during the migration of 67 Meloidogyne javanica juveniles within an Arabidopsis thaliana root.

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Figure 2. 13 Magnified view of the cytoskeletal changes during 68 the migration of Meloidogyne javanica juveniles within an Arabidopsis thaliana root.

Figure 2. 14 An Arabidopsis thaliana root infected with 69 Meloidogyne javanica showing microtubule organisation in root tip cells close to the nematode.

Figure 2. 15 Immunolabelling of a root tip gall in Arabidopsis 71 thaliana infected with Meloidogyne javanica showing the organisation of microtubules in the gall tissue.

Figure 2. 16 Microtubule labelling in a two-day-old gall in an 73 Arabidopsis thaliana root infected with Meloidogyne javanica.

Figure 2. 17 A four-day-old gall in an Arabidopsis thaliana 76 root infected with Meloidogyne javanica immunolabelled for microtubules.

Figure 2. 18 Disruption of xylem in an Arabidopsis thaliana 78 root infected with Meloidogyne javanica.

Figure 2. 19 A 14-day-old gall induced by Meloidogyne 79 javanica in Arabidopsis thaliana shows deterioration of the cytoskeleton in root tissue.

Figure 2. 20 Immunolabelling of a BMM section of an 81 Arabidopsis thaliana infected by Meloidogyne javanica showing the organisation of actin and microtubules in a root gall.

Figure 2. 21 A BMM section of a Meloidogyne javanica- 82 induced Arabidopsis thaliana root gall showed actin and microtubules in a cell.

Figure 2. 22 A GFP-hTalin Arabidopsis thaliana root 24 h after 84 inoculation showing entry by Meloidogyne javanica just behind the root tip.

Figure 2. 23 A GFP-hTalin Arabidopsis thaliana root 48 h after 86 inoculation showed Meloidogyne javanica juveniles migrating within the root.

Figure 2. 24 GFP-hTalin Arabidopsis thaliana showed actin 88 labelling in Meloidogyne javanica feeding sites in a 14-d old root gall.

Figure 2. 25 Immunolabelling of an Arabidopsis thaliana root 90 tip 3 h after inoculation with Pratylenchus thornei shows the organisation of actin and microtubules.

Figure 2. 26 Immunolabelling for actin and microtubules in an 91 Arabidopsis thaliana root 24 h after inoculation with Pratylenchus thornei showed that the cytoskeleton of the root epidermis was modified.

Figure 4.1 Mean number of Meloidogyne javanica observed 132 in roots of tomato, Solanum lycopersicum, grown in sand, sandy clay loam and clay soil three, six and ten days after inoculation.

Figure 4.2 Mean numbers of Pratylenchus thornei observed 134 in roots of tomato, Solanum lycopersicum, grown in sand, sandy clay loam and clay soil three, six and ten days after inoculation.

Figure 5.01 Light micrograph of tomato, Solanum 145 lycopersicum, root shows Meloidogyne javanica second stage juveniles (J2) migrating within 3 days after inoculation.

Figure 5.02 Higher magnification of root tip in Figure 5.01 146

Figure 5.03 Transmitted light micrograph of a Meloidogyne 147 javanica J2 migrating within a tomato, S. lycopersicum, root.

Figure 5.04 Transmitted light micrograph of a tomato, 148 Solanum lycopersicum, root shows several invading Meloidogyne javanica J2.

Figure 5.05 Transmitted light micrograph of a tomato, 149 Solanum lycopersicum, root shows Meloidogyne javanica J2 migrating within the vascular bundle.

Figure 5.06 Transmitted light micrograph of a tomato, 150 Solanum lycopersicum, root shows a Meloidogyne javanica juvenile in a stage between a late J2 and an early J3.

Figure 5.07 Light micrograph of a tomato, Solanum 152 lycopersicum, root shows a Meloidogyne javanica J2 moulting into a J3.

Figure 5.08 Transmitted light confocal image of Arabidopsis 152 thaliana root showing early stage of Meloidogyne javanica J2 moulting.

Figure 5.09 Light micrograph of a tomato, Solanum 153 lycopersicum, root showing a late stage Meloidogyne javanica J3 within root tissue.

Figure 5. 10 Transmitted light micrograph of a Meloidogyne 153 javanica J3 within a tomato, Solanum lycopersicum, root.

Figure 5.11 Transmitted light image of a tomato, Solanum 155 lycopersicum, root infected by Meloidogyne javanica.

Figure 5.12 A tomato, Solanum lycopersicum, root with an 156 adult Meloidogyne javanica within.

Figure 5.13 An adult Meloidogyne javanica within a tomato, 157 Solanum lycopersicum, root.

Figure 5.14 Tomato, Solanum lycopersicum, root infected by 159 Pratylenchus thornei. Figure 5.15 A tomato, Solanum lycopersicum, root observed 160 six days after inoculation with Pratylenchus thornei.

Figure 5.16 Pratylenchus thornei infecting a tomato, Solanum 161 lycopersicum, root.

Figure 5.17 Surface tissue of tomato, Solanum lycopersicum, 162 root damaged due to infection by Pratylenchus thornei.

Figure 5.18 Pratylenchus thornei infecting a tomato, Solanum 163 lycopersicum, root.

Figure 5.19 Pratylenchus thornei within tomato, Solanum 164 lycopersicum, roots.

ABBREVIATIONS a Actin af actin filaments ab actin bundles acf alternating cell files of root hair forming and non root hair forming cells asn actin surrounding nucleus awc actin outside a wounded cell BMM butyl methyl methacrylates BSA bovine serum albumin C Chromosomes c Cortex cb shed cuticle of nematode body ce damage localised to a single cell cg cortical tissue forming the gall clp cells which have lost their polarity col co-localised actin and microtubules ct tail region of the shed cuticle cw cell wall cx Cortex DAPI 4’,6-diamidino-2-phenylindole dc damaged/disrupted cortical tissue DMSO dimethyl sulphoxide EGTA ethylene glycol bis(2-aminoethyl ether)-N,N,N’N’-tetraacetic acid f Fluorescence G, g Gall gc giant cell GFP green fluorescent protein I Interphase lr lateral root M Metaphase Mb median bulb MBS m-Maleimidobenzoyl-N-Hydroxysuccinimide ester MgSO4 magnesium sulphate mn/nm migrating nematode mt Microtubules N Nucleus n Nematode NaCl sodium chloride Na2HPO4 disodium hydrogen phosphate nb nematode body nb1 nematodes within vascular bundle ne nematode entry nh nematode head ni nematode intestine nn nematode neck nt nematode tail

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nv nematode breaching vascular bundle Omt obliquely arranged microtubules P Prophase PBS a buffer solution containing NaCl, Na2HPO4 and KH2PO4 ph phragmoplast PIPES Piperazine 1, 4-bis (2-ethanesulfonic acid) PME a buffer containing PIPES, EGTA and MgSO4 PMSF Phenylmethylsulphonylfluoride ppb preprophase band PSEs proto sieve elements rh root hairs Rmt randomly arranged microtubules sm secretory material sp Spindle st swollen root tissue st1 unidentified structures st2 nematode stylet T Telophase Tmt transversely arranged microtubules tn turning nematode vb vascular bundle w wound caused by nematode Wmt wavy microtubules x normal xylem xd/dx disrupted xylem vessels xf xylem around feeding site

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SUMMARY

Plant parasitic nematodes infect almost all crop plants and annually cause losses of millions of dollars worldwide. The relationship between these pathogens and their hosts is still poorly understood in spite of several decades of research. In this research project, I attempted to investigate different aspects of this host-parasite relationship with respect to root-parasitic nematodes, and the effect these nematodes have on growth and development of the affected plants.

I studied the host-parasite interactions using two different host plants, Arabidopsis thaliana and Solanum lycopersicum, and two nematodes which contrasted in their modes of infection, life cycle and pathogenicity: the root-knot nematode, Meloidogyne javanica, and the root lesion nematode, Pratylenchus thornei. Studies on A. thaliana were conducted in the laboratory under aseptic conditions in Petri dishes, and studies on tomato were conducted in the greenhouse as pot culture experiments.

In my first investigation I examined the changes occurring in the cytoskeleton of A. thaliana root cells due to infection by M. javanica and P. thornei. This experiment was conducted over a period of one year from March 2003 to March 2004 at the Research School of Biological Sciences, Australian National University, Canberra, ACT in the laboratory of Drs Geoff Wasteneys and David Collings. The plant cytoskeleton plays a vital role in almost all cellular functions including cell signalling, cell division and wound response. I used three different investigative techniques to compare the effects of plant parasitic nematodes on the host root cytoskeleton. These techniques were whole root double immunolabelling, double immunolabelling of BMM sections and use of GFP-hTalin-transformed A. thaliana. Samples were observed using confocal laser scanning microscopy.

I found all three techniques to be effective in studying cytoskeletal changes in infected roots. Whole root immunolabelling and GFP-hTalin-transformed A. thaliana

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were most useful for studying the early stages of infection. However, immunolabelling of BMM sections was most effective in the later stages of infection to study large and thick gall tissue, as the transmission of light through thick tissue was not adequate to obtain high quality images by the other two techniques.

Whole root immunolabelling showed that both M. javanica and P. thornei initially (even within the first 3 h) caused similar changes in A. thaliana. All stages of M. javanica were observed in A. thaliana roots; their migration and development was observed for 14 d. However, observations on P. thornei could not be conducted beyond the first 24 h after inoculation due to bacterial contamination. While P. thornei damaged A. thaliana roots, they were not observed within roots at any stage during this observation period. This was probably because the small diameter of the roots made them unattractive for entry by this larger nematode species.

Material labelled for actin was observed to accumulate in a rounded or disc-shaped plug on the root surface and appeared to line the wound surface on the inside of the cell within the first 24 h after inoculation. Microtubules maintained their orientation, giving support to the cell wall. Increased actin and microtubule labelling was detected in nematode-infected root tissue with all three techniques used. In A. thaliana roots inoculated with M. javanica, this was especially prominent during initial entry and feeding site formation but was not as noticeable during nematode migration. This may have been due to a change in permeability of the cell walls resulting from nematode infection. A diffuse fluorescence was observed whenever actin fixation was not adequate. Once the M. javanica nematode entered the root, it followed a path (probably associated with physiological or chemical signals derived from the different cell types) during different stages of its migration to reach its ultimate feeding site. Microtubules in host cells close to the nematode body in its migration path were in a wavy, rather than a taut, arrangement. In nematode feeding sites, giant cells were formed around the head of the nematode and numerous small cells were observed towards the posterior of the nematode, forming the gall. Unusual divisions were observed in nematode-infected root tissues, with abnormal spindles and incomplete phragmoplasts in a region where cell divisions do not normally occur. Using immunolabelled BMM sections I observed well-defined microtubules and actin bundles in an infected cell. Giant cells appeared to have lost their growth

polarity and anisotropy. These giant cells were spindle-shaped, with two tapering ends and with an enlarged middle section, indicating that there may be significant differences between the end (cross) walls of cells and the walls parallel to the longitudinal axis of the root.

I subsequently conducted a temporal investigation in tomato, as an adjunct to a pot trial, to determine the similarities and differences in the way in which the two nematode species entered, migrated and caused damage, as well as their development within roots. My observations of roots were made using light microscopy and staining with acid fuchsin. This study showed, in detail, the entry and migration for both nematode species and, for M. javanica, feeding site formation and developmental life stages. This enabled longer time for observations of P. thornei than was the case with the A. thaliana study. The initial stages of infection were consistent with the Arabidopsis results. However, I was able to observe the later stages of infection, which were markedly different between the two nematode species, although at a lower magnification than in the confocal study.

To investigate the effects of the nematode-plant interactions that I had observed in my laboratory studies on plant growth and development, I conducted a pot trial using tomato seedlings, which ran over a period of 11 weeks, using the two previously- studied nematode species. Treatments involved inoculation of pots with 5000 larvae. Different pots were infested weekly, from the date of planting to one week before harvest. At harvest (11 weeks), I recorded the length of aerial shoots, the number of green leaves, dry weight of shoots and roots, the combined number of flowers and flower buds and the number, fresh weight and diameter of fruits. M. javanica significantly (p < 0.05) reduced shoot length, the diameter of the largest fruit, and dry weight of shoots and roots, while P. thornei significantly (p < 0.05) reduced shoot length, number of green leaves, the combined number of flowers and buds, and the dry weight of shoots and roots. While fruit from plants infected with M. javanica had similar moisture content to those in the untreated control, fruit from plants infected with P. thornei had lower moisture content than comparative control plants. These results indicate that the differences between the modes of infection of these two nematode species that I observed in the laboratory studies are manifested in host plant growth and development.

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Texture, together with pore size and sufficient moisture, is reported to be the primary soil factor influencing plant root-attacking nematode movement towards host roots for infection. I investigated the movement of M. javanica and P. thornei through soils towards the roots of tomato, S. lycopersicum, in a pot trial. For this I used three soils with different textures; namely sand, sandy clay loam and clay.

While only a small proportion (< 19% in M. javanica and < 11.5% in P. thornei) of the original inoculum of 300 nematodes were recorded in the tomato roots, this is consistent with previously published studies. However, contrary to previous reports, there were many fewer nematodes present in roots of plants in sandy clay loam soil than in clay soils. The likely explanation is that the sandy clay loam soil had substantially high organic matter, electrical conductivity and silt. This indicates that soil chemistry and, probably, soil biology can play an even more important role than texture in determining nematode movement to and their damage of plant roots.

I conclude that M. javanica and P. thornei differ in the way they interact with root tissues during the later stages of infection, although the host cytoskeletal reaction is similar during initial entry by both nematodes. These effects are manifested in the subsequent growth and development of host plants. A combination of soil physical, chemical and biological factors influence the movement of nematodes through soil to roots of host plants, and, in the case of P. thornei, may also influence length of time they spend within roots.

The results of my study provide opportunities for further in-depth examination of the infection process, to understand the host-parasite interaction between nematodes and their host plants. A better understanding of the nature of these interactions will assist in formulating improved control strategies for these important nematode pests.

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CHAPTER 1

General Introduction

Nematodes are unsegmented, bilaterally symmetrical round worms which belong to the phylum Nematoda. The word nematode is made up of two Greek root words, nema- (thread) and -oides (like) (Decker and Sveshnikova, 1989). Their body is protected by a strong outer cuticular layer which has transverse and/or longitudinal striations. They have successfully evolved to adapt to all living conditions from the frigid polar regions to dry deserts, from valleys to mountaintops, and even salt and fresh water (Weischer and Brown, 2000). The abundance of nematodes on Earth has been explained eloquently by Cobb (1914) thus:

“In short, if all the matter in the universe except the nematodes were swept away, our world would still be dimly recognizable, and if, as disembodied spirits, we could then investigate it, we should find its mountains, hills, vales, rivers, lakes and oceans represented by a thin film of nematodes. The location of towns would be decipherable, since for every massing of human beings there would be a corresponding massing of certain nematodes. Trees would still stand in ghostly rows representing our streets and highways. The location of the various plants and animals would still be decipherable, and, had we sufficient knowledge, in many cases even their species could be determined by an examination of their erstwhile nematode parasites.”

Nematodes vary in length from less than 100 µm (marine nematodes) to several metres (sperm whale parasites). Some species of nematodes can spend their whole life in the soil feeding on soil microorganisms and organic matter (free-living), while some need other living organisms such as animals or plants to complete their life cycle (parasitic). Parasitic nematodes utilise animals (including humans) or plants as

hosts. Parasitism can cause diseases and even death of hosts, in severe cases. Animal parasites include roundworm, whipworm, heartworm, Toxoplasma and filarial worm (Maggenti, 1981).

Not all nematodes are harmful. Some species of insect parasitic nematodes (entomopathogenic nematodes) of genera Steinernema and Heterorhabditis have been used for biological pest control in agriculture (Wouts, 1991; Nguyen and Hunt, 2007).

Nematodes have also benefited the scientific research community. It is notable that the first multicellular organism to have its genome completely mapped was the free- living nematode Caenorhabditis elegans (Ankeny, 2001). This nematode is considered a “model organism” for research into various cellular functions relating to animals, including cell differentiation and meiosis. In November 2005, scientists from National Aeronautics and Space Administration (NASA) reported that canisters containing live C. elegans had been recovered from the wreckage of the spaceship Columbia. These nematodes had been part of an international experiment on board the ill-fated ship and their recovery was ample proof that life can indeed survive the searing heat of their passage through the earth’s atmosphere on re-entry (Szewczyk and McLamb, 2005).

There are a number of species of plant parasitic nematodes. A few of them are highly significant economically as they cause severe damage to plants. Some of the most important plant parasitic species which occur in crop fields worldwide include Meloidogyne spp. (root-knot nematodes), Heterodera spp. (cyst nematodes), Pratylenchus spp. (lesion nematodes), Ditylenchus spp. (stem and bulb nematodes), and Rotylenchulus spp. (reniform nematodes) (Decker and Sveshnikova, 1989; Nickle, 1991).

Tainter and Baker (1996) listed nematodes that cause diseases of almost 2000 other crop plant species as well as forest trees. Meloidogyne spp. and Pratylenchus spp. were found on elm, cherry and other hardwood trees as endoparasites (i.e. they enter, feed and reproduce within host roots), while other species including Xiphinema spp., Hoplolaimus spp., and Bursaphelenchus spp. are ectoparasitic (i.e. they remain

outside of roots and penetrate with only a small portion of their bodies where they feed on the epidermal and cortical cells; sometimes they may enter wholly into the cortical tissue).

Some nematodes have developed strategies for highly successful parasitism while others have not. Evolutionary adaptation by Meloidogyne has ensured its survival by its ability to induce specialised feeding structures within the host plant. It also produces highly environmentally-resistant eggs covered in a protective gelatinous matrix which aids survival in adverse soil conditions.

Plant parasitic nematodes are of significant economic importance in agriculture because they cause billions of dollars in crop losses annually, world wide (Webster, 1972; Southey, 1978; Wyss, 1992). Growth and yield suppression of host plants occurs mainly because of the large quantities of nutrients used by the nematodes for their growth and reproduction; the reduction in host root growth due to infection by nematodes also affects absorption of adequate water and nutrients from the soil (Hussey, 1985).

Above ground symptoms of severe nematode infection are usually similar to plants with root damage; infected plants show yellowed and falling leaves, stunted growth and low yield. The infected plants wilt temporarily under conditions of water stress. In severe cases, plants die in large patches in the field. Below the ground, the symptoms on host roots caused by different species of nematodes can vary considerably (Hunt et al., 2005). Their mode of feeding can range widely, from destructive external feeding on roots to the formation of highly specialised feeding sites induced within the host, which nourish the parasite until it completes its life cycle.

1.1 Choosing nematodes for the study

1.1.1 Economic importance of Meloidogyne spp. and Pratylenchus spp. in crop production

The root-knot nematodes, Meloidogyne spp., and the root lesion nematodes, Pratylenchus spp., are two of three most important root parasitic nematode genera occurring worldwide. Both of these nematode genera infect many species of cultivated and wild plants (Fielding, 1959; Nickle, 1991; Luc et al., 2005). They damage plants as a result of their parasitism and reduce yields, resulting in significant economic losses to farmers (Eisenback and Triantaphyllou, 1991; Luc et al., 2005).

The root-knot nematode, Meloidogyne spp., is as widespread in Australia as it is around the world. In a survey conducted in Victoria (Harris, 1984) Meloidogyne was found in 35% of vegetable crops sampled and was the most common nematode. Meloidogyne was also found in 53% of vineyard samples. The root lesion nematode, Pratylenchus spp., was found in 40% of samples from fruit crops and 18% from vineyards. Extensive yield losses caused by these nematodes have also been reported in fruit crops such as banana (De Waele and Davide, 1998), pineapple (Stirling and Kopittke, 2000), sugarcane (Cadet and Spaull, 2005), vegetable crops such as carrot (Hay, 2000), and potato (Stirling and Wachtel, 1985); and flower crops such as Boronia (Webb, 2005).

In the rice growing regions of South East Asian countries M. incognita has caused poor seedling establishment (Bridge et al., 2005). Grain yield was also reduced by very high population densities of M. javanica; yields decreased by 40% when 8000 eggs and juveniles /dm3 of soil were present at sowing (Babatola, 1984). Damage was more severe under upland conditions (Fademi, 1984). In flooded rice, the tolerance limit of seedlings in nurseries is one J2 (second stage larva) of M. graminicola per cm3 of soil (Plowright and Bridge, 1990). In South East Asian upland rice, Meloidogyne combined with Pratylenchus poses a serious threat economically (Prot et al., 1996). Both Meloidogyne and Pratylenchus infect citrus

and other tropical and subtropical fruit crops (Duncan, 2005; El-Borai and Duncan, 2005).

Pratylenchus thornei was found to cause 27 to 70% yield loss in wheat crops in South Australia (Taylor and McKay, 1993; Nicol et al., 1999), and crop losses have been reported to be up to 50%, in New South Wales (Doyle et al., 1987) and 50-85% in Queensland (Thompson et al., 1997). In citrus, growth reduction of up to 49-80% was noted (O’Bannon and Tomerlin, 1973) due to Pratylenchus coffeae infection. Approximately 30% yield reduction was recorded in rice (Prasad and Rao, 1978; Plowright et al., 1990) in fields infected with Pratylenchus.

Important root and tuber crops like potato, cassava, sweet potato, beetroot, and those grown regionally such as yams, taro and coleus are also severely affected by both Meloidogyne and Pratylenchus (Bridge et al., 2005; Scurrah et al., 2005; Vovlas et al., 2005). In the case of tuber and root crops the storage quality and marketability is also severely reduced. Results of a field experiment conducted in 1988 at the College of Agriculture, Vellayani, India showed that almost 100% of Coleus parviflorus tubers harvested from Meloidogyne-infected plots were unmarketable after storage due to spoilage (Pazhavarical 1988, unpublished). Meloidogyne also causes severe damage to the root system of peanut crops, and up to 100% losses have been reported (Dickson and DeWaele, 2005).

In addition to acting as a nutrient sink, these pathogens exacerbate host damage by providing entry points for other secondary microorganisms such as fungal, bacterial and viral plant pathogens (Fielding, 1959).

A large number of survey studies have showed the occurrence of Meloidogyne and Pratylenchus in the root zone of crop plants; however far fewer have actually quantified the economic losses incurred from infection by these pathogens. This could be because it is difficult to accurately separate crop losses caused by these nematodes under field conditions from crop losses caused by other factors such as the effect of other plant pathogens, adverse environmental conditions and the tolerance or resistance of the host species (Ferris, 1981). The present study aims to

minimise these variables by employing a pot culture experiment using only the host (tomato) and the selected pathogens, M. javanica and P. thornei.

Although above-ground symptoms of infection by Meloidogyne and Pratylenchus are similar, these two species of plant parasitic nematodes have highly contrasting biology, life cycles and modes of parasitism. Even though they cause heavy and widespread losses in crops, very little is known about their host-parasite interactions, particularly during the early infection stages. The lack of detailed information concerning the post-infection stages of the pathology of these nematodes was the motivating reason for choosing these nematodes for this study. This research, thus, attempted to investigate the early post-infection phases of M. javanica and P. thornei in the host plants.

The study consisted of four parts. In the first part, A. thaliana was used as a model host to study the cytoskeletal changes within the root cells during the early stages of infection. Tomato, Solanum lycopersicum L., was used as host in the second part to study the movement of the nematodes in different soil types. The third part of the project dealt with the effect of nematode infection of tomatoes at different phenological stages on plant growth and yield, and also used tomato as the host. In the fourth part I observed the early stages if infection of M. javanica and P. thornei on tomato

1.1.1.1 The root-knot nematode Meloidogyne javanica (Treub, 1885) Chitwood, 1949

Meloidogyne javanica is one of the most successful plant parasites capable of infecting more than 2000 plant species (Hussey et al., 1994). They pose a significant threat to plant production worldwide (Sasser, 1980). Much of the morphological, taxonomic and other research work on this nematode was done in the late 1800s and in the 1900s.

M. javanica has been widely recognised by farmers for centuries because of the easily noticeable, typical galls it produces on the plant roots. According to Decker and Sveshnikova (1989), the first official written report of root-knot nematodes was

by the Reverend Miles Joseph Berkeley in 1855, a clergyman who noted galls on cucumber roots in England. Since that time there have been several more reports and many attempts at its nomenclature. Among the various synonyms used for Meloidogyne spp. are Caconema (Cobb, 1924) Hypsoperine (Karssen, 2002) and Heterodera radicicola (Muller, 1884). In 1887, Meloidogyne exigua, which caused a root-knot on coffee, was identified and described by Goeldi in Brazil. In 1932, Goodey reclassified Heterodera radicicola as Heterodera marioni (Karssen, 2002). However, it was Chitwood (1949) who removed them both from the genus Heterodera as he found that they were different from cyst nematodes. As the oldest name existing at the time was Meloidogyne, that name was given to the nematode.

The word Meloidogyne is derived from combining two Greek words, Meloido- (apple-shaped) and -gyne (female). Originally, identification was based on perineal patterns (a finger-print like pattern found on the cuticular surface of females, posteriorly). More recently, differential host range tests (Taylor and Sasser, 1978), chromosome counts (Triantaphyllou, 1966), juvenile head structure (Eisenback and Hirschmann, 1981) and analysis of various proteins, including isozymes produced by the nematode, have been used for its taxonomy (Eisenback and Triantaphyllou, 1991).

Meloidogyne is the most economically important genus of plant parasitic nematodes, and occurs worldwide; in Australia, Asia, USA, Europe, Africa and South America (Luc et al., 2005). It consists of about 60 species. Of these, only four species, M. incognita (Kofoid and White, 1919) Chitwood, 1949, M. javanica, M. arenaria (Neal, 1889) Chitwood, 1949 and M. hapla Chitwood, 1949, cause most of the crop damage and economic losses worldwide. Meloidogyne spp. have a host range of up to 2000 crop species. However, some species of nematodes are specific to a single plant host e.g. Meloidogyne pini whose host range is restricted to Pinus spp. (Jepson, 1987).

The International Meloidogyne Project

Considering the significant economic damage Meloidogyne spp. cause in crops worldwide and the fascinating and complex nature of their host-parasite interactions

an International Meloidogyne Project was set up by USAID, from 1975 to 1985, with the following objectives:

1 Identify species and biotypes of the root-knot nematode within eight geographical regions 2 Determine resistance / susceptibility of food crops 3 Identify sources of resistant germplasm 4 Evaluate crop responses in each region 5 Study the variability within species, host reaction, morphology, cytogenetics etc. 6 Evaluate environmental factors influencing distribution in each area 7 Develop integrated crop protection systems for control of root knot nematode in each region.

The organisations which took part in this project included the International Potato Center (CIP) Peru, the International Crops Research Institute for the Semi-Arid Tropics (ICRISAT) India, International Institute of Tropical Agriculture (IITA) Nigeria, the International Center for Tropical Agriculture (CIAT) Columbia, the International Center for Maize and Wheat Improvement (CIMMYT) Mexico, and the Asian Vegetable Research and Development Center (AVRDC) Taiwan. Through the work of this project there was increased awareness of the importance of Meloidogyne in developing countries. The distribution, frequency and relative importance of the different nematode races in agricultural crops and soils was surveyed. New species were discovered, described, and identified, based on more reliable characteristics. Phyletic relationships were clarified on the basis of cytogenetic and biochemical characteristics. Ecological factors affecting survival, distribution and pathogenicity were described. In addition, research capabilities of developing nations were expanded through conferences, publications, and field and laboratory training sessions. A comprehensive treatise covering all aspects of Meloidogyne studied through this project was published in a two-volume book (Sasser and Carter, 1985; Barker et al, 1985).

M. javanica was used for studies in my project. This nematode has a host range of about 800 host plants. It occurs within the latitude range of 33o N to 33o S and prefers warm, dry climate and areas with a prolonged dry season. It is scarcely found

in cold areas where the temperature drops below 3o C (Eisenback and Triantaphyllou, 1991). Optimum temperature range for growth and multiplication is from 25-30o C (Luc et al. 2005).

Morphology and lifecycle

Morphology and life cycle of the various species of Meloidogyne are similar within the genus. They survive in the soil as environmentally-resistant eggs. When a suitable host is available they hatch into motile, vermiform, second stage juveniles (J2) and actively seek host tissue in response to, as yet unknown, stimuli. The J2 stage of Meloidogyne spp. can vary between 290-912 µm in length; body length of M. javanica J2 is between 402-560 µm.

When they reach the adult stage, they show sexual dimorphism. Adult males remain vermiform, from 700-2000 µm in length. A distinguishing characteristic is that their body is twisted 180o along its length. Adult females become swollen and spherical, their body size ranging from 0.44-1.3 mm long and 0.325 to 0.7 mm wide, with their body enclosed within the root tissue and the head and neck protruding anteriorly (Eisenback and Triantaphyllou, 1991). They are pearly white in colour, smaller than a pin head and are just visible to the naked eye. Their oesophagus has a large and prominent muscular median bulb. M. javanica reproduce by mitotic parthenogenesis. Under conditions of environmental stress such as overcrowding, unfavourable host and high temperature, sex reversal occurs with more females turning into males and when favourable conditions return, more males turn into females (Bird, 1971). This is a highly effective survival strategy for the nematode.

J2 are the only infective stage in the life cycle of the nematode (Noe, 2003). J2 can probably survive in a quiescent stage in the soil for long periods. As they have to depend on food stores in the body until a feeding site is established in a suitable host, infectivity decreases with depletion of these reserves (van Gundy et al., 1967). J2 locate their host by the help of the cephalic sensory organs composed of six inner labial sensillae and two amphids but the exact mechanism is unknown. Although their preferred infection sites are in the zone of elongation behind the root tips, they are also attracted to apical meristems, penetration sites of other J2, root galls, cut

surfaces of roots and the site of lateral root rupture (Decker and Sveshnikova, 1989; Krusberg and Nielsen, 1958; Bird, 1959). Actively elongating roots are more attractive to J2 than slow growing roots (Godfrey and Oliveira, 1932).

The nematodes breached the epidermal cells of the host root by constantly rubbing their anterior mouth parts against the root surface and by stylet thrustings. Video microscopic studies on cultured A. thaliana (Wyss et al., 1992) showed that Meloidogyne used its stylet to rupture the epidermal cells to gain access to the host. It then migrated apoplastically (i.e. between cells) towards the root tip. Endo and Wergin (1973) reported that cells were separated along the middle lamella during migration. Cells along the path of migration were distended and compressed but did not show any signs of feeding. These cells along the migratory path showed increased cytoplasmic density (Jones and Payne, 1978). At the tip, the absence of a differentiated endodermis allowed the nematodes to turn around and enter the vascular cylinder (Niebel et al., 1993). Then they migrated towards the differentiating region, where they became stationary in a suitable feeding spot. Once the feeding spot was selected, each J2 produced highly specialised feeding sites called “giant cells” and fed from them (Jones, 1981a; Wyss, 1992; Wyss et al., 1992). If the J2 failed to initiate giant cells within the root, it either starved to death or migrated out of the root to locate another suitable host root. Galls were formed around the feeding nematode by hyperplasia of cells adjacent to the body of the nematode. During its lifetime the J2 underwent three moults, increasing its body diameter each time, and at the same time retracting the body anteriorly, finally attaining the globular form of the adult female after the third moult (Karssen, 2002). Typically, the adult female was located with its head, embedded in the vascular region among the giant cells and its body directed outwards. On maturity, the female laid an egg mass containing up to 500 eggs into a gelatinous matrix secreted by the rectal glands towards the outer surface of the root. This gelatinous matrix putatively suggested pectolytic, cellulolytic and proteolytic activity (Orion and Franck, 1990) and its function was to protect and preserve the eggs during adverse environmental conditions. The females died within the host root after oviposition, and the egg masses within the matrix existed in soil until favourable conditions returned. Multiple infections by M. javanica on root tips caused enlargement and suppression of growth of the root (Godfrey and Oliveira 1932).

Stages of normal mitotic cell division in plants

It is pertinent here to explain about how the cell divides in plant cells. Normal mitosis in plant root cells consists of five main stages:

(1) Interphase This is the resting phase of the cell in-between divisions. (2) Prophase This phase is characterised by the formation of spindles and the shortening and thickening of chromosomes. (3) Metaphase In this phase the chromosomes get attached to the spindle and are arranged in the plane of the equator of the spindle, called the metaphase plate. (4) Anaphase In this phase the chromosome pairs separate and move to the opposite poles of the cell. (5) Telophase During this phase two daughter nuclei are formed and the mother cell divides into two daughter cells by the formation of a phragmoplast which lays down the new cell wall.

The plant cytoskeleton has an important and dynamic role in cell division. Microtubules are the major component of the preprophase band, spindles and phragmoplast. Actin has also a major role to play in cell division, and is found to co- localise with microtubules in the phragmoplast. Functions of the plant cytoskeleton are reviewed in detail, in relation to uninfected and nematode infected cells of the root in Chapter 2.

Giant cells induced by Meloidogyne in the host

After a period of migration, during which it follows a defined path to the differentiating region, Meloidogyne becomes stationary and induces a few cells around its head to form greatly enlarged “giant cells”. These giant cells provide food

to the nematode for the remaining period of its life. One of the earliest experiments conducted by Bird (1961) showed that Meloidogyne was alone responsible for giant cell production, and that an unidentified plant-growth promoting substance, absent in uninfected root tissue, was present in the root galls. In this experiment the nematode was killed with a needle, resulting in subsequent necrosis and vacuolation of the giant cell. This was accepted as proof of a continuous stimulation from the nematode and that the continued presence of the female was required for the maintenance of the giant cell. Alternatively, it was suggested that the metabolic drain that the plant is subjected to by the nematode stimulated the plant to generate signals that are responsible for maintaining the giant cell (Jones, 1981b). The nematodes can survive without formation of a gall but cannot live in the absence of a functioning giant cell (Webster, 1969).

Cytoplasm of giant cells appear very similar to that of active meristematic cells with numerous cell organelles such as golgi bodies, endoplasmic reticulum, mitochondria and a large number of small vesicles accumulated around partially formed cell plates between two newly divided cells (Jones and Payne, 1978). The physiology of giant cells and their similarity to meristematic cells has been reviewed by Jones (1981a).

Genes in giant cell formation

Genes control all aspects of plant development. Therefore, it follows that the ability of a nematodes to establish and maintain a highly specialised feeding site within the host allows it to manipulate the normal genetic expression of the host to suit its parasitic requirements. On the other hand, in nematode-resistant host plants, the host may react to the parasite by switching on defense responses or hypersensitive responses such as early cell senescence and programmed cell death. Several genes have been identified in plants following nematode infection, which relate to susceptibility, resistance or other defence-related responses e.g. hypersensitive response and programmed cell death.

Researchers have followed the expression of genes induced during early response to M. incognita infection in both susceptible and resistant alfalfa, Medicago sativa, and also in different plant organs. Based on these analyses, the genes that are induced

early in nematode infection are related either to metabolic pathways or to stress/defense (Potenza et al., 2001). In spite of this, the apparent absence of host wound or expression of defence response is an extremely interesting characteristic of the initial stages in a compatible root knot nematode-host interaction. Copious secretions from the second stage larvae (J2) stylets during migration may actively suppress host responses. This suppression may also explain the increased susceptibility of nematode galls to invasion by secondary pathogens. Some promoters seem to be induced in the host while the giant cells and nematodes are enlarging but not in the early stages of infection when J2 are still migrating in the root (Hansen et al., 1996).

Using promoter-gusA constructs, it was found that in A. thaliana (ecotype C24) the expression of a large number of genes was influenced during the development of nematode feeding structures (Goddijn et al., 1993). Genes upregulated in giant cells included the RB7 in tobacco (Wilson et al., 1994). Extensin genes were induced by M. javanica on tobacco (van der Eycken et al., 1996). High extensin gene expression was observed during the whole second larval stage (a two-week long phase of establishment of the feeding site) of the nematode. During later stages of this interaction, expression gradually decreased. Extensin gene expression was found in at least three different tissues of the gall (Niebel et al., 1993).

Meristematic cells in plants are usually found in the growing tips, for example, shoot and root tips. These cells are usually characterised by high metabolic activity, large number of small vacuoles, numerous organelles and a dense cytoplasm. These characteristics are also found in Meloidogyne-induced giant cells (Jones, 1976; Jones and Payne, 1978) indicating that the nematode may have found ways to induce meristematic characteristics in the host to create and maintain giant cells. Class 1 knotted gene (KNOX) and PHANTASTICA (PHAN) expression required for normal meristem maintenance (Reiser et al., 2000; Timmermans et al., 1999) was specifically expressed in tomato giant cells (Koltai and Bird, 2000).

Origin of the giant cell

Even though giant cells are known to be feeding structures induced by Meloidogyne and are known to be absolutely essential for the survival of the nematode, the exact mechanism of the formation of these structures is not fully understood. But, as similar giant cells are induced in a wide variety of host plants, the process that leads to the formation of giant cells may involve some fundamental and widely conserved aspects of plant biology (Niebel et al., 1996; Bird and Bird, 2001).

Giant cell progenitors, or initials, have been identified to form from parenchymatous cells adjacent to the xylem (Krusberg and Nielsen, 1958; Jones and Payne, 1978), but other preferred sites have been observed including cells from pericycle, cortex and epidermis (Bird, 1996). Irrespective of their origin, induction of giant cells starts within 24 h after infection. Each root-knot nematode triggers the development of up to five to seven cells to develop into permanent nurse cells or “giant cells” (Sijmons et al., 1994). Giant cells, as their name suggests, are cells that are enlarged several- fold compared to normal cells, sometimes reaching dimensions of 600 µm in length and 200 µm in diameter (Jones, 1981b). The cell walls of the giant cells are thickened irregularly and have extensive wall ingrowths. They have many mitochondria, and may contain as many as 100 nuclei that have undergone endo- reduplication (Wiggers et al., 1990). As the giant cells enlarge in size they show abnormal nuclear division. The first nuclear division does not result in a difference in size of the neighbouring normal parenchymatous cells. Cell division is not completed with each nuclear division and they become multinucleate (Huang and Maggenti, 1969; Jones and Payne, 1978). After telophase, the partially formed cell plate becomes attached to the mother cell wall and forms a stub or flange. Electron microscopic studies (Jones and Payne, 1978) showed vesicles, golgi bodies, endoplasmic reticulum, mitochondria and small vacuoles accumulated around partially formed cell plates between two daughter nuclei. Nuclei are highly variable in size and shape, irregularly lobed and amoeboid (Huang and Maggenti, 1969). Linkages between neighbouring nuclei by lobes are commonly observed (Starr, 1993; Wiggers et al., 1990). Several studies have reported synchronous nuclear division in different host plants (Krusberg and Nielsen, 1958; Bird, 1961; Owens and Specht, 1964; Smith and Mai, 1965 [cited by von Mende, 1997]); Huang and

Maggenti, 1969; Jones and Payne, 1978). Abnormal spindle orientation occurs, resulting in irregular mitosis. The cytoplasm of giant cells resembles that of active meristematic cells with abundant cell organelles like golgi bodies, mitochondria, ribosomes, endoplasmic reticulum and polysomes (Jones and Northcote, 1972; Jones and Dropkin, 1975; Jones and Gunning, 1976; Jones and Payne, 1977) and several small vacuoles (Jones and Payne, 1978; Wergin and Orion, 1981). Growth of the giant cells and their DNA content reach a peak just before the nematodes start laying eggs (2-3 weeks after infection). After this time the size of giant cells and the DNA content of their nuclei decline significantly (Bird, 1972). This might be the stage in infection that puts the most strain on the plant, due to the induced growth of feeding structures and galls and the large quantity of nutrients withdrawn by the nematode at the time.

Function of giant cells

The principal function of giant cells is to unload the downwardly translocating photosynthate from the phloem and present it in a form accessible to the parasite (Bird. 1996). Even though giant cells are symplastically isolated from neighbouring cells (Jones and Dropkin, 1976; Böckenhoff and Grundler, 1994), the phloem unloading, which occurs, may involve pump-driven transmembrane channels (Bird, 1996). Movement of solutes has been reported to be unidirectional from phloem to syncytium and may be apoplastic. Anomalous loading of carboxyfluorescein and radioactive carbon from sieve element companion cell complexes occur specifically into the syncytium (Dorhout et al., 1991; 1993; Böckenhoff and Grundler, 1994; Böckenhoff et al., 1996). In A. thaliana infected by the cyst nematode Heterodera schachtii (Schmidt, 1871), the fluorescent dye, lucifer yellow, injected into the syncytium (the nematode feeding structure located within the stele of the root) stayed in the structure, showing that it is symplastically isolated from the surrounding root tissue (Böckenhoff and Grundler, 1994). From the syncytium, both fluorescent and radioactive labels were withdrawn by feeding nematodes. Studies revealed that Heterodera infection triggered the formation of special type of phloem cells called “unloading phloem” that allowed macromolecular trafficking of green fluorescent protein into syncytia (Hoth et al., 2005). The nematode induced formation of phloem containing an approximately three-fold excess of sieve elements over companion

cells. This newly formed phloem exhibited typical properties of unloading phloem, similar to those described in other sink tissues. A similar mechanism could be induced in hosts by Meloidogyne, which would make food translocation into the giant cells easier.

Giant cells and transfer cells

The giant cells induced by Meloidogyne on plants have been found to have similar characteristics as ‘transfer cells’ normally found in plants. The endosperm in Arabidopsis and Lepidium virginium contains large, multinucleate cells. They also exhibit extensive wall ingrowths and lack of larger organelles, suggesting an important role in loading of maternal resources into the developing seed (Nguyen et al., 2000). Similar to the structure of the transfer cell walls these giant cells have wall ingrowths at regions of high solute transfer. Wall ingrowths increase the surface area of the cell wall many-fold, thereby enhancing transport of solutes across the plasma membrane (Jones, 1976; Gunning, 1977; Gunning and Steer, 1996; Offler et al., 2002). Reactivation of the cell cycle in specific plant cells such as the endosperm finally results in these transfer cells. Some similar process may be taking place in the development of nematode-induced feeding sites.

A study using cryofixation of Arabidopsis indicated that no mitotic phragmoplasts are present in syncytial endosperm cells, but rather mini-phragmoplasts are formed (Otegui and Staehelin, 2000). Several mini-phragmoplasts appear to participate in making a single plate by independently initiating vesicle fusion events at several locations simultaneously (Verma, 2001). Fragmented phragmoplasts were also observed in giant cells caused by Meloidogyne infection (De Almeida Engler et al., 2004).

Role of nematode secretions in nematode entry and migration in plants

A plant cell wall normally contains cellulose, hemicellulose, pectin and structural protein (Green, 1964; Cosgrove, 1993; 1997; 1999).

Cellulose is an essential component of the plant cell wall. This suggests the involvement of cellulase for facilitating initial infection by nematodes (Keen and Roberts, 1998) as well as other enzymes, without which the infection process is made difficult. In the case of root-knot nematodes these secretions may originate in the hypodermis (Roze et al., 2008) oesophageal glands, the chemosensory amphids or the cuticle (Reviewed in Caillaud et al., 2008). The nematode cuticle, or glycocalyx, exudates from secretory organs and stylet exudates are variously believed to play important roles in nematode migration within the host root and also in induction of giant cells (Bird, 1992). Within plants, nematode movement putatively involves wall- degrading enzymes produced by the nematodes, which may be up-regulated by nematode feeding. Several enzymes have been isolated and characterised from nematode-infected cells including -glucuronidase (Barthels et al., 1997), -1, 4- endoglucanase (Rosso et al., 1999), other family 5-endoglucanases (Bera-Maillet et al., 2000) and pectin acetylesterase (Vercauteren et al., 2002). Studies with infective stages of M. incognita showed the presence of two genes encoding pectate lyase and poly galacturonase. The transcription was localised to the pharyngeal gland of the nematode suggesting that they could be the components of stylet secretions. These enzymes may help parasitism by decomposing the compounds of the cell wall of the host plant root (Davis et al., 2000), creating suitable conditions for nematode migration within the plant root. M. javanica chorismate mutase 1, an oesophageal gland protein, putatively altered plant cell development allowing nematodes to establish a parasitic relationship with the host plant (Doyle and Lambert, 2003). Calreticulin has also been reported to suppress plant defence responses (Jaubert et al., 2005). However, there is still insufficient information available to create a comprehensive model, primarily because the sequences of several of these genes have no similar sequence found in the existing databases. The functional analysis of putative parasitism genes will, therefore, be a challenge for the future (Vanholme et al., 2004).

Factors affecting cell enlargement in plants following Meloidogyne infection

In a normal plant cell, enlargement is regulated cell-specifically. Cell expansion is regulated by several factors, including water, light, temperature and gravity.

Chemical factors such as gibberellins, auxins and other plant hormones also affect cell expansion, as well as cell wall mechanics, hydraulics and biochemical processes.

Cytokinins are compounds that promote cytokinesis or cell division. Cytokinins have been implicated in formation of lateral roots and nodules in legumes. A study of the feeding structures induced by nematodes in Arabidopsis found that processes involved in formation of nematode feeding structure and lateral root development share common anatomical features and also common patterns of genetic expression. It has been observed that lateral roots are induced very near where galls are induced by Meloidogyne (Sasser, 1954). A high level of expression of the gene ARR5 (a cytokinin responsive gene) was induced when J2 Meloidogyne reached the differentiating vascular bundle in Lotus japonicus roots, and also during early stages of the nematode-plant interaction. ARR5 expression was specifically absent in mature giant cells, although dividing cells around the giant cells continued to express this reporter. The same pattern was observed using a GFP reporter driven by the ARR5 promoter in tomato (Lohar et al., 2004). Genes and plant growth hormones affected by nematode infection were reviewed in detail by Bird and Koltai (2000). It is significant that normal cytokinesis is absent in giant cells (Huang and Maggenti, 1969) although close to the posterior body of the nematode, host cells divide prolifically to form the characteristic gall. Areas of gall formation have been noted to be prolific in lateral root formation.

1.1.1.2 The lesion nematode Pratylenchus thornei (Sher and Allen, 1953)

Pratylenchus was first identified by De Man in 1880 from a meadow in England as Tylenchus pratensis (Decker and Sveshnikova, 1989). This genus was named from a combination of a Latin word pratum (meadow), and two Greek words tylos (knob) and enchos (spear). Similar to Meloidogyne, this genus has also been known under several synonyms in its taxonomic history. As in the case of Meloidogyne, initial taxonomy was based on morphological characteristics (Loof, 1991) but, of late, new methods such as restriction fragment length polymorphism (RFLP) and isoelectrofocusing (ISF) (André et al., 2001) have been successfully used for classification. There are about 70 described species, of which at least ten species cause widespread economic damage in about 400 host crop plants including cereals, fruit trees, ornamentals and turf in Australia, USA, India, Europe and South Africa (Loof, 1991). Pratylenchus is only third behind Meloidogyne spp. and Heterodera spp. in economic damage caused to crops worldwide.

Morphology and life cycle

This nematode can lay eggs either in soil or within the host plant. Eggs hatch into second stage juveniles which are vermiform. Juveniles and adults in the genus resemble each other and differ only in length. Body length of an adult Pratylenchus can vary from 0.3 to 0.9 mm and index a (body length/body height) is 20-30 (Loof, 1991).

Pratylenchus thornei, used for the studies in my project, is a serious pest of wheat in Australia and USA, but also infects many other crops including other cereals, legumes and banana all over the world (Loof, 1991; Luc et al., 2005). It mainly occurs in heavy soils of Mediterranean (moderate) climatic regions from 20° C to 30° C. However, it can also occur in other types of soil. Length of the life cycle varies depending on the species and temperature; from 30-40 d at 25° C - 30° C to 50-60 days at 20° C. The body of P. thornei is about 0.45-0.77 mm long and comparatively stouter than average, with index a of 26-38 (Loof, 1991).

P. thornei is a migratory nematode which can spend its entire life cycle within the plant (endoparasitic) or outside it (ectoparasitic) (Sijmons et al., 1994). This nematode also has the ability to live part of its life within the root and then migrate out of the root to live in soil. When host plants are not available, Pratylenchus can survive in soil as a free-living nematode.

Kurppa and Vrain (1985) recorded the process of initial infection by Pratylenchus in detail in strawberry plants. They observed that the nematodes migrated to the root hair zone and selected an epidermal cell by rubbing the cell surfaces with their labia and stylet. Stylet thrusting, salivation, predigestion and ingestion phases were recognised during the feeding process. Penetration occurred after the nematodes bored a series of holes in the cell wall and forced their way in. Penetration also occurred, sometimes through root hairs. Sometimes the nematodes withdrew and opened another hole on the opposite side of the hair. Pratylenchus were not usually seen infecting cells in the root tip or cell elongation zone. In other hosts, Pratylenchus aggregated just behind root tips (Bird, 1960) but no penetration of the root cap was observed. Following local exploration of cell surfaces the area beside an intercellular wall was pierced, and the stylet thrusted several times into the cell. Similar observations have been made in vegetable crops. Histopathological studies of chickpea, Cicer arietinum, genotypes infected by Pratylenchus thornei showed that they always migrated through epidermal and cortical cells by breaking down cell walls along the path of migration. Occasional damage to endodermal cells was observed (Castillo et al., 1998), but the nematodes mostly confined their parasitism to cortical cells (Zunke, 1990). Traces of feeding on epidermal cells (i.e. minute holes made through the cell walls) were difficult to recognise with certainty in SEM or semi-thin sections. Damaged cells collapsed and died rapidly. Root cells thus penetrated by the nematode formed an entry point for secondary infection by other pathogenic microorganisms, finally resulting in necrosis of the roots. Nematodes were observed to move through epidermal cell layers to the cortical layers centripetally. Many eggs were laid in the cortex (Kurppa and Vrain, 1985). Ectoparasitic feeding on root hairs of host plants was also observed (Zunke, 1990).

In some cases, these nematodes fed on the surface epidermal cells and remained ectoparasitic, not penetrating into the root cortex. Most of them moved along the root

surface, feeding at several different sites. These feeding sites later showed as small necrotic patches or lesions (Webb, 1990). When conditions become unfavourable in the root, the nematode migrated to the soil and survived in a free-living form until favourable conditions returned.

1.1.2 Control of plant parasitic nematodes

Cultural practices

Crop rotation with alternating host and non-host plants and using cover crops which produce chemicals toxic to nematodes can reduce their populations. Hot water treatment of seed tubers and incorporation of organic matter are also effective control methods. In warm climates, flooding the field and turning over the top soil to expose lower layers to the sun can also achieve reduction in their population. These methods are labour intensive and need extensive planning for maximum effectiveness (Sikora et al., 2005).

A few nematode-resistant crops have been developed over the past few decades. Very few crop plants have been found to be resistant to all four major species of Meloidogyne, because of their wide host range. The Mi gene, however, confers resistance to all four species. Rootstocks resistant to M. javanica have been developed for stone fruit such as almonds, peaches, plums and nectarines, fruit crops such as grape vines and citrus; vegetables such as tomato, carrot, potato, soybean, cowpea and capsicum, as well as field crops such as cotton, lucerne, groundnut, tobacco and wheat (Hussey and Janssen, 2002). In trials conducted by the International Meloidogyne Project from 1975 to 1985, only 1.3% of crop plants showed high resistance to all nematode populations. Main crop plants tested were tomato, potato, chickpea, pigeon pea, bean, Desmodium, wheat, barley and corn (Sasser and Carter, 1985). Resistance genes sometimes become ineffective at high temperatures (Starr et al., 2002). In some cases resistance to one race of M. incognita can be broken by another race (Riggs, 1991; Starr et al, 2002). Breeding for resistance and crop rotation may not be highly effective for control of Pratylenchus because of its cosmopolitan nature and wide host range (De Waele and Davide, 1998; De Waele and Elsen, 2002). Some root stocks resistant to P. thornei include

varieties in peach, plum, apple, almond and rose. Pratylenchus-resistant varieties of cereals, root crops, banana, lucerne, strawberry have also been reported (De Waele and Elsen, 2002). The search for new plant genes useful in developing nematode resistance in crops is continuing through research.

Recently, molecular techniques have been used to develop resistance to nematodes in host plants. A PCR-based marker which was tightly linked to the root knot resistance gene Mi was developed in tomato (Williamson et al., 1994), and quantitative trait loci (QTLs) associated with resistance in wheat to P. thornei and P. neglectus were located and tagged (Zwart et al., 2005). Nematode resistance genes have been reviewed in detail by Williamson and Kumar (2006).

Nematicides

Nematicides are often effective and reduce economic losses caused by nematodes, especially in combination with cultural practices. However, the toxicity of these chemicals towards non-target organisms, and the environmental and health risks they pose to humans (Starr et al., 2002) and the environment have caused concern over their continued use. Recently, there has been increasing awareness of the risks of nematicides which, in most cases, outweigh their benefits (Thomason, 1987). Many of them are extremely potent nerve toxins and cause not only immediate effects following exposure, but in a number of cases, delayed effects. Costa (2005) has comprehensively reviewed these and other current issues with regard to organophosphate toxicity in non-target species.

Biological control

Organisms such as fungi, bacteria and nematodes naturally occurring in the soil can compete with or cause diseases in plant parasitic nematodes and can, thus, be used for their control. This topic has been reviewed by several researchers over the past 40 years (Jatala, 1986; Sikora, 1992; Cook, 1993). This method may be less effective than nematicides, due to the large quantity of inoculum necessary to achieve successful results as well as the inability to target specific pathogens (Mankau, 1981).

Integrated Pest Management

An integrated approach, involving combinations of resistant cultivars, crop rotation, biocontrol agents, and clean and correct cultural practices as well as judicious use of nematicides is likely to be the most effective in controlling plant parasitic nematodes

(Barker and Koenning, 1998). Adopting a combination of two or more control methods such as crop rotation, cultural practices and crop sanitation is the most effective way to control both Meloidogyne and Pratylenchus. The main objective of this approach is to keep the population of plant parasitic nematodes below economic threshold levels (Bird, 1987; Sikora et al., 2005).

1.1.3 Use of molecular techniques and phylogenetics in nematode identification

Several molecular techniques have been developed to identify plant parasitic nematodes. Pratylenchus spp. was rapidly and readily identified using a reverse dot blot test (Uehara et al., 1999). Although Powers et al. (2005) found that a nucleotide sequence from the highly conserved 18S region did not discriminate between M. arenaria, M. incognita and M. javanica, Qiu et al. (2006) subsequently used polymerase chain reaction (PCR) to identify these species. Sequence characterised amplified region (SCAR) based PCR assays have also been used to identify M. incognita, M. javanica and M. arenaria (Zulstra et al., 2000) as well as P. thornei (Carrasco-Ballesteros et al., 2007). Real-time PCR has been used used to identify P. penetrans (Sato et al., 2007), and P. zeae, M. javanica, and the dagger nematode Xiphinema elongatum (Berry et al., 2008).

Subbotin et al. (2008) conducted phylogenetic analyses of Pratylenchus spp., revealing at least six distinct major clades which were generally congruent with those defined by characters derived from lip patterns, number of lip annules, and spermatheca shape. The molecular taxonomy and phylogenetics of Meloidogyne spp. has been recently reviewed by Adams et al. (2009). There exists scope for further analysis and study in the field of phylogenetics based on these studies.

Studies of the genome and gene transcripts of some nematode species have also provided valuable information with regard to gene identification and expression. For example, expressed sequence tags (ESTs) were conducted on Pratylenchus spp. and putative nematode-specific and Pratylenchus-specific genes were identified (Mitreva et al., 2004). A genome-wide overview of gene expression during the interaction between M. incognita and Arabidopsis (Jammes et al., 2005) suggested that there is a suppression of plant defence associated with nematode feeding site formation. Studies using ESTs of M. hapla and M. incognita have been completed recently (Bird et al., 2009). Such studies will aid the further understanding of nematode-host interactions.

1.2 Choosing model host plants for the study

A. thaliana was chosen for cytoskeletal studies in my research program because it has several attributes that would assist in understanding various aspects of the host- pathogen relationship, especially at the cellular level (Dolan et al., 1993; Bowman, 1994). Arabidopsis has a relatively simple root system. Its small size and cell number make it an ideal system for studying infection by microorganisms such as nematodes. This also makes it amenable to a variety of experimental manipulations. Individual cells can be observed in real time and their development recorded. It has a simple and well-studied root anatomy, and the cell lines have been mapped. Cell files in the primary root are relatively constant in number. This may help in determining the origin of feeding site formation in roots infected by Meloidogyne.

Moreover, Arabidopsis has the ability to grow well under controlled laboratory conditions. It can grow well at normal room temperatures (20-25° C), both in vitro and in the greenhouse. It has a small seed size and is extremely prolific, and has a rapid generation time (~5 to 6 weeks under optimum conditions [Bowman, 1994]). These characteristics enable large populations to be established quickly and easily maintained in the laboratory. A. thaliana has been widely recognized as a model plant for studying plant-nematode interactions (Sijmons et al., 1991; Niebel et al., 1994). The thin, translucent roots enable observation of early stages of the infection process. In my investigations it served as a suitable model to study the interaction between plant roots and nematodes in vitro.

Tomato, S. lycopersicum, was chosen for the nematode population dynamics and movement studies because it has been known to be a good host for both Pratylenchus and Meloidogyne in pot culture and in Petri dishes. It is easy to grow from seed in the laboratory and in the greenhouse over a range of temperatures.

1.3 Aim and Objectives

1.3.1 Aim of this study

The main aim of this project was to understand the plant-nematode interaction in roots during the post-infection period. The work was composed of four different studies: 1. The changes in the cytoskeleton in root cells of A. thaliana due to infection by Meloidogyne javanica and Pratylenchus thornei (Chapter 2) 2. The effect of the nematodes Meloidogyne javanica and Pratylenchus thornei on growth and yield of tomato plants (Chapter 3) 3. The movement of Meloidogyne javanica and Pratylenchus thornei in different types of soil such as sandy, loamy and heavy clay soils (Chapter 4) 4. Plant infection by Meloidogyne javanica and Pratylenchus thornei, and their migration and development in tomato roots (Chapter 5).

1.3.2 Rationale behind the aim and objectives of this study

The ability of Meloidogyne during post-infection stage to induce re-differentiation of root cells into feeding sites is a fascinating aspect of its biology (Gheysen et al., 1996). Although there is detailed published information on the infection process of both Meloidogyne and Pratylenchus, there is a paucity of information about the early post-infection phase at the cellular level, especially in relation to cytoskeletal changes in the host. This is fundamental to understanding the processes involved in disease incidence and severity. Nematode infection can affect the normal cycle of cell division in roots, especially in the case of infection by Meloidogyne where giant cells are induced by the nematodes in the plant at the site of nematode feeding. Pratylenchus can affect the cytoskeleton due to their destructive feeding on root cells. The aim of my first study was to elucidate the early effects of parasitism by Meloidogyne and Pratylenchus on the cytoskeleton of root cells of Arabidopsis, with special reference to microtubules and actin microfilaments.

The above-ground symptoms caused by Meloidogyne and Pratylenchus can be similar. However, these nematodes exhibit significantly contrasting epidemiology,

including life cycles and modes of parasitism. The differences between them in regard to their modes of infection and the paucity of information available on comparative crop losses caused by them were the motivating reasons for choosing these nematodes for my second study on tomato.

Meloidogyne and Pratylenchus spend the majority of their life in soil, and may even move over large distances to reach host plant roots. The physical and chemical properties of soil affect nematode movement towards the host root zone, and thus, the level and intensity of infection in the field. There is very little information on the comparative effect of different types of soil on nematode movement and root infection. Therefore, the purpose of my third study was to determine the influence of soil texture, using sandy, sandy loam and clay soils, on the movement of the nematodes Meloidogyne and Pratylenchus to roots of tomato and subsequent levels of infection.

As previously stated, Meloidogyne and Pratylenchus have contrasting epidemiology, including life cycles and modes of parasitism. Tomato is a natural host for both of these species, and it has a larger and more complex root system than Arabidopsis. This is likely to provide opportunities for longer-term observations on the post-entry aspects of nematode infection. Therefore, the purpose of my fourth study was to conduct temporal observations of nematode infection, migration and development within host plant roots.

CHAPTER 2

Changes in the cytoskeleton in root cells of Arabidopsis thaliana due to infection by Meloidogyne javanica and Pratylenchus thornei

2.1 Introduction: The cytoskeleton in animals and plants

The cytoskeleton in plant cells consists of two filamentous structures, actin microfilaments (also referred to as F-actin) and microtubules, while most animal cells contain these and a third array of intermediate filaments. Actin microfilaments are made up of polymerised globular protein subunits (G-actin) whereas microtubules are composed of polymerised dimers of two distinct but related globular proteins α- and β-tubulin. Free actin and tubulin dimers are found in the cytoplasm, and these proteins have been highly conserved during eukaryotic evolution (Alberts et al., 2002).

The actin microfilament is made up of actin monomers and is about 8 nm in diameter (Figure 2.01). The head to tail arrangement of actin monomers gives the actin microfilament a structural polarity. Two parallel protofilaments are twisted around each other in a right-handed helical pattern. Actin microfilaments have a fast- growing plus end and a slow-growing minus end; actin is bound to adenosine triphosphate (ATP) which is hydrolysed to adenosine diphosphate (ADP) after polymerization, which enables dynamic instability and treadmilling (Alberts et al., 2002).

Figure 2.01 The structure of actin microfilaments

(A) A schematic representation of actin sub-units shows the plus end and minus end. The yellow part represents ATP (ADP when bound in polymerisation). (B) Diagrammatic representation in which several actin sub-units similar to those in (A) have polymerised to form an actin microfilament. The microfilament also shows plus and minus ends. (C) Electron microscopic image of an actin microfilament. [From: Alberts et al. (2002) Molecular Biology of the Cell, 4th Ed.]

Figure 2.02 The structure of microtubules

(A) Diagrammatic representation of a microtubule shows a dimer composed of one -tubulin (light green) and one -tubulin (dark green) subunit. The red represents GTP, which in -tubulin hydrolyses to GDP during polymerisation. The GTP in -tubulin is not hydrolysed. (B) Several dimers polymerise to form a microtubule protofilament which, similar to the dimer. The plus end bestows the microtubule with the property of dynamic instability and treadmilling which is used in forming the various microtubular structures required for cell functions. (C) A microtubule formed by the bonding of thirteen protofilaments similar to that shown in (B). Both (B) and (C) exhibit a plus end and a minus end. (D) Electron microscopic image of a microtubule. [From: Alberts et al. (2002) Molecular Biology of the Cell, 4th Ed.]

A microtubule is constructed as a relatively stiff hollow tube, about 25 nm in external diameter, composed of 13 protofilaments arranged parallel to each other (Figure 2.02). Each protofilament is made up of heterodimers composed in turn of one - and one - tubulin monomer. The pattern of longitudinal - and lateral - bonds is repeated in a helical manner along the length of the microtubule. The microtubule has a structural polarity because of the arrangement of the subunits. As with actin microfilaments, each - and - tubulin monomer is bound to a nucleotide, in this case a guanosine triphosphate (GTP) molecule. The GTP bound to the -tubulin monomer

cannot be hydrolysed or exchanged. On the other hand, the GTP bound to a -tubulin can be hydrolysed to a guanosine diphosphate (GDP) form and is easily exchangeable when dimers are released from the polymer by disassembly. This property of the microtubule plays an important part in microtubule dynamics. The nucleation of a microtubule is from a -tubulin ring complex which acts as a base for - and -tubulin to assemble into a microtubule. As with microfilaments, the two ends of a single microtubule polymerise at different rates. The end which grows and shrinks fast is called the plus end and the end which grows and shrinks at a slower rate is called the minus end (Alberts et al., 2002).

The cytoskeleton functions with the help of certain accessory proteins that link these structures to other cell components, as well as to each other. Those associated with actin are called actin-binding proteins (ABPs) whereas those binding to microtubules are microtubule-associated proteins (MAPs). ABPs and MAPs include motor proteins which move organelles and other cargo along the cytoskeleton. Motor proteins associated with microtubules are dynein and kinesin, although dyneins do not exist in higher plants (Lawrence et al., 2004) whereas myosin is the motor associated with actin microfilaments.

Although the G-actin and tubulin proteins that make up the cytoskeleton have been highly conserved during eukaryotic evolution, and while ABPs and MAPs have also been conserved although not so strongly, there are significant differences between the animal and plant cytoskeletons. In animal cells, cytoplasmic microtubules generally radiate from the centre of the cell towards the cell periphery. In plants, interphase microtubules are dispersed all over the cortex reflecting diffuse growth. In animal cells, there is a well defined microtubule organising centre (MTOC), called the centrosome, located near the nucleus. Microtubules are nucleated from numerous -tubulin ring complexes located in the pericentriolar material of the centrosome, with their minus end associated with the centrosome such that they radiate in an astral or star-like form with their plus ends towards the cell periphery. In higher plants, well-defined MTOCs have not been identified (Azimzadeh et al., 2001; reviewed in Wasteneys and Galway, 2003; Wasteneys and Yang, 2004). Instead, plant microtubules are nucleated at various sites distributed all around the nuclear

envelope (Alberts et al., 2002) and in dispersed sites in the cell cortex (Dixit and Cyr, 2004).

In animal cells, actin nucleation occurs at the plasma membrane, catalysed by complexes of proteins called actin-related protein complexes (ARP complexes) because two of their constituent proteins are ARP2 and ARP3. Similar processes have been shown to occur in plants. A member of the formin family of proteins, AFH1, has been identified recently as a probable actin nucleator in A. thaliana (Michelot et. al., 2005). Several ARPs have been identified in plants that are involved in specific cellular functions (Kandasamy et al., 2004). Some perform cytoplasmic functions such as actin organisation, cell morphogenesis and cell polarity. Some are localised to the nucleus and are involved in protein transcription, chromatin organisation and DNA repair. Actin plays an important role in plant cell growth in conjunction with several actin binding proteins (reviewed in Hussey et al., 2006).

Both -tubulin and ARPs, responsible for nucleating microtubules and actin respectively, are evolutionarily ancient and tightly conserved among a wide variety of eukaryotic species. Plant and animal actins share 85% of amino acid sequence identity (Alberts et. al., 2002). However, ABPs are less conserved between animals and plants (Gardiner and Marc 2003; Hussey et al., 2002), which may account for the differences between the cytoskeletons of animal and plant cells. Some ABPs, like plant profilin, are less than 50% similar to those in animals, and some ABPs identified in animals such as zyxin are not seen in plants (Alberts et al., 2002).

2.1.1 Functions of the plant cytoskeleton

All major plant cell processes, including cell division, growth and enlargement, involve the active participation of the cytoskeleton (Smith, 2002). The plant cytoskeleton plays an important role in guiding growth polarity in plant cells. However, microtubules and actin differ in the type of functions they carry out in the cell (reviewed in Wasteneys, 2000).

2.1.1.1 The functions of plant actin

Actin filaments act as tracks for organelle movement, facilitate cytoplasmic streaming, anchor chloroplasts in response to favourable light conditions and respond to infection by other organisms by rearranging themselves (Alberts et al., 2002). As subcellular trafficking in higher plants is actin-specific, the transport machinery of plant cells differs considerably from that in animal cells where much intracellular transport is generated by microtubules. Actin plays important roles in plant morphogenesis and cell signalling, especially in association with guard cell movements, mechano- and gravity- sensing, plant host-pathogen interactions and wound healing (Volkmann and Baluška, 1999).

2.1.1.2 The functions of plant microtubules

In addition to the functions of the cytoskeleton mentioned earlier, microtubules function in cell wall deposition and the formation of the preprophase band, spindle fibres and phragmoplast during cell division (Dashek and Harrison, 2006). Microtubules also play a role in gravitropic bending (Himmelspach et al., 1999; Lloyd et al., 2000) and have also been linked to cell vacuolation and rapid cell elongation (Baluška et. al., 1992). In the meristematic and elongating region of the root, arrangement of microtubules is mainly transverse. In the differentiating region of roots, microtubules are arranged obliquely or longitudinally. During cell division microtubules are found in different structural formations including pre-prophase bands, spindles and phragmoplasts. In order to achieve the remodeling required to form these different structures, tubulin subunits constantly polymerize and depolymerize following cues from the cell.

Cell elongation and maintenance of growth polarity is under the control of microtubules (Wasteneys, 2000; Mathur and Hülskamp 2002). The direction in which plant tissue cells expand reflects in the alignment of microtubules in the cortical array (Yuan et al., 1994). Cells have one predominant microtubule alignment (transverse, oblique or longitudinal). Re-orientation probably involves movement of stable or dynamic microtubules. When microtubules and co-aligned wall microfibrils

are arranged transversely around the cell, turgor pressure is channeled into cell elongation. However, various agents (such as wounding, ethylene, abscisic acid) can cause the microtubules to reorient by 90° so that they become aligned parallel to the cells’ long axis, allowing lateral expansion instead of elongation. The mechanism by which microtubules undergo rapid shifts of alignment is crucial to understanding growth control in plants.

2.1.2 Dynamics and organisation of the microtubules within plant cells

One of the aims of this study is to try to find if any microtubular responses are induced in root cells by nematode infection. It is well-known that microtubules in plant cells change orientation in response to cues from within and without the cell.

By micro-injecting pea (Pisum sativum) epidermal cells with rhodamine-conjugated brain tubulin and optically sectioning them by confocal laser scanning microscopy, a study followed labelled microtubules for up to 2 h as they reoriented. Reorientation did not occur by complete depolymerisation of microtubules in one orientation followed by polymerization of a new array in another orientation. Instead, increased numbers of discordant microtubules in non-transverse alignment appeared in particular locations. Neighbouring microtubules then adopted the new alignment, so that there was a stage during which different alignments co-existed before the array on the outer tangential cell phase finally adopted a uniform, steeply oblique/longitudinal configuration. Rapid fluorescence recovery after photo bleaching confirmed that bundles of cortical microtubules are not stable, but exhibit properties consistent with dynamic instability. The study concluded that dynamic microtubules offer a mechanism for rapid growth responses to a range of physiological stimuli (Wymer et al., 1997).

Close resemblance has been noted between cellulose microfibril orientation and microtubule orientation (Hogetsu, 1986; Hogetsu and Oshima, 1986; Abe et al., 1995; Sugimoto et al., 2000; Baskin et al., 2004; Emons et al., 2007). In the elongating region of the root, microfibrils were deposited transversely to the root axis. In the region where elongation ceased, obliquely oriented microfibrils discontinuously overlaid the transverse ones. The dynamic nature of microtubules is

further illustrated during cotton fibre development. During fibre initiation and early elongation, microtubules have a generally random orientation. Microtubules re-orient into shallow pitched helices as elongation and primary wall deposition continue and into steeply pitched helices during secondary wall deposition. Accompanying the changes in orientation are increases in microtubule length, number, proximity to plasmalemma and a decreased variability in orientation of the microtubules (Seagull, 1992).

2.1.3 Interaction between actin and microtubules in plant cells

Microtubules and actin microfilaments interact in plant cells (Collings and Allen, 2000; Collings, 2008). During cell division, microtubules and actin work together by co-aligning in laying down the phragmoplast, which is required for constructing the cell plate that eventually forms a new cell wall between daughter nuclei during cell division (Wasteneys and Collings, 2004). Actin and microtubules co-align and function during axial growth of cells, cell elongation and control of the plant shape (Collings and Allen, 2000; Staiger and Hussey, 2004).

Fine, transversely oriented cortical actin filaments have been observed in all cells of the elongation zone, including the epidermis, cortex and vascular tissue (Blancaflor, 2000). The orientation of cortical actin shifts formed a predominantly transverse orientation to oblique, longitudinal or transverse and/or random arrangements as cells mature. The reorientation of cortical actin in maturing root cells mimics the behaviour of cortical microtubules reported in other studies. Results of this study by Blancaflor (2000) suggested that cortical microtubules and actin microfilaments can respond in a coordinated way to environmental signals.

In most plant cells, cortical actin comprises a dynamic array of single actin microfilaments and/or small bundles that interact with the plasma membrane. These interactions may function to anchor the entire actin cytoskeleton, and contribute to cell signalling. In cells elongating by diffuse growth, cortical actin is typically transversely oriented, parallel to microtubules (Sonobe and Shibaoka 1989; Blancaflor, 2000; Collings and Wasteneys, 2005). Drug studies demonstrate that cortical actin is necessary for the precise rearrangements of cortical microtubules that

control cell expansion and plant morphogenesis (Takesue and Shibaoka, 1998). However, in other cases, cortical actin depends on the cortical microtubules for its organization. This suggests that the cortical cytoskeleton is a closely controlled system involving feedback between actin and microtubules (Collings and Allen, 2000).

Although they may have somewhat different functions in the plant cell, the fact that microtubules and actin microfilaments co-localize and function together point to the need to study them both during nematode infection.

2.1.4 Changes in the plant cytoskeleton due to external factors

When plants interact with their environment, they have to respond to different stimuli. These include changes in sunlight, external pressure, and interactions with pathogenic and non-pathogenic organisms and wounding. During these interactions, extensive signalling occurs in plant cells, and in the case of interactions with other organisms, signalling occurs between these organisms and the host plant. The dynamic cytoskeleton plays an active role in these responses (Wasteneys and Yang, 2004). Extensive cytoskeletal rearrangements occur in plant hosts following wounding and infection by pathogenic and non-pathogenic micro-organisms. These interactions between the host and parasite may start during the first few minutes of association (Heath 1997; 2000) and presumably continue during all stages of infection.

Pathogenic fungi

Studies indicate that infections of plants by pathogenic fungi cause changes in the cytoskeleton. These changes may be associated with changes in growth or the plant’s response to the pathogen. For example, granular deposits of reaction material form when Botrytis allii attempts to penetrate onion leaves, resulting in a striking polarization of actin microfilaments that form a network of filaments focussed towards the penetration site (McLusky et al., 1999). Similarly, the rice blast fungus Magnaporthe grisea can induce actin rearrangement during infection of rice (Xu et al., 1998).

Microtubules can also reorient during fungal infections. In flax leaves infected by the rust fungus, Melampsora lini, microtubules were observed to form a transverse array in the mesophyll cells. Microfilaments radiated through the cytoplasm from the nucleus. In an incompatible reaction, microtubules and microfilaments were extensively reorganised in the mesophyll cells in contact with fungal hyphae before penetration of these cells by the pathogen. After the initiation of haustoria, microtubules disappeared from the infected cells and growth of haustoria ceased. In compatible interactions, reorganisation of the cytoskeleton is much less frequent (Kobayashi et al., 1994). Microtubules have been reported to also be affected following infection of cowpea, Vigna unguiculata, by the rust fungus, Uromyces vignae. In two resistant cultivars, observations of microtubule organization prior to cell death suggested a possibility of multiple pathways for cellular degradation during the hypersensitive response (Škalamera and Heath, 1998).

Fungal symbiosis

The host cytoskeleton can be affected by fungal symbiosis with a close association existing between the cytoskeleton and invading symbiotic fungal hyphae in orchids (Uetake et al., 1997) and Scots pine (Niini et al., 1996). Cytoskeletal re-organization was also observed in root cells of Medicago truncatula during development of an arbuscular mycorrhizal symbiosis with Glomus versiforme. Extensive remodelling of the microtubule cytoskeleton occurred from the early stages of arbuscule development until arbuscule collapse and senescence. Cells adjacent to infected cells also showed reorganisation of microtubules (Blancaflor et al., 2001).

By contrast, infection by a mycorrhizal fungus in Lotus japonicus (wild type) epidermal cells resulted in only limited cytoskeletal changes during the early stages of fungal infection. Later, however, the growing hypha was surrounded by microtubules and microfilaments, while the host cell nucleus moved close to the fungal hypha. In contrast, symbiosis-defective mutants responded with disorganisation and disassembly of microtubules and microfilaments before and during fungal penetration. Due to the close relationship between host cytoskeleton organisation and compatibility with the fungus, it appears that certain proteins are necessary for correct re-organisation of the epidermal cell cytoskeleton in the

presence of the fungus and for avoiding hypersensitivity-like reactions (Genre and Bonfante, 2002).

Bacterial symbiosis

As in the case of fungal symbiosis, root nodules formed in plants in response to infection by bacterial symbionts can affect the cytoskeleton. These symbiotic bacteria activate mitotic activity in cortical cells giving rise to the nodule primordium (Cárdenas et al., 1998) through secretion of Nodulation factor (Nod factor). Nod factors change the host cell organisation by acting on the actin cytoskeleton (Lhuissier et al., 2001). Bean, Phaseolus vulgaris, root hairs infected by Rhizobium etli were studied by microinjecting with fluorescein isothiocyanate-phalloidin (Cárdenas et al., 1998). They concluded that Nod factors alter the organization of actin microfilaments in root hair cells, which could be a prelude to the formation of infection threads. A study of nodulation in alfalfa roots infected by rhizobia showed that microtubules in bacteria-free alfalfa nodules were not disorganized while those in bacteria-infected nodules were. This indicated that microtubule disorganization required the presence of bacteria (Timmers et al., 1998).

Wounding

Changes in the cytoskeleton occur after a plant cell is wounded. In Nitella internodal cells, regeneration of actin filament bundles was studied in order to identify the mechanisms by which microtubules are oriented (Foissner and Wasteneys, 1999). In the different types of wounds investigated, subcortical actin bundles regenerated parallel to the direction of cytoplasmic streaming. Microtubule orientation patterns, however, varied according to the nature of wound formation and the type of wound wall eventually produced. These results indicated that microtubules regenerated in wounds are merely co-aligned with actin filament bundles because of passive alignment through hydrodynamic forces created by cytoplasmic flow. Wounding was followed by a dramatic shift in ionic currents in Pisum sativum roots (Hush and Overall, 1989). Cortical actin microfilaments reorient around a wedge-shaped wound in pea to lie in planes parallel to the contours of the wound (Hush and Overall, 1992). However, this rearrangement occurred later in time than a similar microtubule

rearrangement, which was 2-5 h post-wounding (Hush et al., 1990). Dynamic rearrangement of microtubules occurred in the absence of actin microfilaments to change cell polarity in response to wounding (Hush and Overall, 1992).

2.1.5 Immunofluorescence and the study of plant cytoskeleton

The plant cytoskeleton has been studied using immunofluorescence techniques for many decades (reviewed in Lloyd, 1987). In recent years, several advances have been made in the immunofluorescent visualization of the cytoskeleton in plants, and in Arabidopsis in particular. The principle of immunolabelling consists of first locating a protein in a sample by using the specific (primary) antibody, which binds to the specific protein. Then a fluorescent secondary antibody is used to label the primary antibody. When the secondary antibody is illuminated it emits fluorescence through which we can visualize the selected protein. Most antibodies to visualise actin and microtubules were developed from animal antigens, but the conservation of actin and tubulin proteins between animals and plants means that the antibodies recognize plant proteins (Hush et al., 1991; Collings et al., 1995).

The advent of the confocal laser scanning microscope has also been a major advance in imaging the plant cytoskeleton because of the clearer images obtained by this method (Hepler and Gunning, 1998). Confocal microscopy uses point illumination with lasers and a pinhole to generate an in-focus only image. All out of focus light is eliminated. Computer software then uses the collected optical sections to generate 2D or 3D images. These visualizations help understand the observed structures (Sheppard and Shotton, 1997).

An alternative method for immunolabelling thicker plant tissue involves embedding, sectioning and then immunolabelling the sections (Baskin et al., 1992). This method has the advantage of observing structures deeper within tissue that conventional laser illumination systems on most confocal microscopes cannot penetrate.

Collings and Wasteneys (2005) described a novel method to visualize both actin and microtubules concurrently. After a refined fixation step, they treated the sample with

two primary antibodies concurrently followed by two secondary antibodies for actin and microtubules respectively. This protocol required confocal microscopy (see below).

2.1.6 GFP in the study of plant cytoskeleton

Green fluorescent protein (GFP), isolated from the jellyfish Aequorea victoria, is a naturally fluorescent, non-toxic protein, which fluoresces in response to UV or blue light. At first it was used widely as a marker in animal cells where it was used to label individual, living cells in vivo so that their development could be followed without harmful manipulation such as dye microinjection or immunohistochemistry (Butner and Kirschner, 1991; Brand, 1995; Ludin and Matus, 1998). GFP has been widely used to study plant anatomy and physiology (Chalfie et al., 1994; Pang et al., 1996).

Several studies have been conducted in living cells of A. thaliana expressing GFP fusion proteins to determine microtubule organisation and dynamics (Haseloff et al., 1998; Ueda et al., 1999). In Arabidopsis cell cultures, microtubules of the preprophase band, spindle and phragmoplast were clearly visible in sequence within 0-10, 20-30 and 40- 50 minutes after the start of cell division (Hasezawa et al., 2000) and similar studies have been conducted in tobacco cell cultures (Kumagai et al., 2001). Comparison of these processes with non-transgenic GFP TUA6 Arabidopsis showed similar timings for the stages of cell division, showing that the normal cell cycle processes were not significantly affected by GFP (Hasezawa et al., 2000).

GFP fusion proteins have also been used to study the actin cytoskeleton in plants. GFP-mTalin, a fusion of GFP to the actin-binding domain of the mouse ABP talin, can serve as a non-invasive marker for the actin cytoskeleton in a number of different cell types of Arabidopsis (Kost et al., 1998). Subsequently, GFP-hTalin (a fusion to the homologous domain in human talin) has also been proved as a probe for microfilaments in Arabidopsis (Takemoto et al., 2003). These probes are not, however, without problems.

In a comparison study, GFP-hTalin-expressing Arabidopsis plants were found to express roughly twice the total GFP fluorescence of another microfilament labelling probe, GFP-fABD2 (a fusion protein between GFP and the second actin-binding domain of A. thaliana fimbrin) (Sheahan et al., 2004). However, GFP-hTalin seemed to cause artificial aggregation of actin networks, suggesting that talin fusions may not accurately depict actin organisation and turnover in plant cells. Further, seedlings transformed with GFP-mTalin showed retarded growth in hypocotyls, inflorescences, siliques and leaves compared to wild type or GFP-fABD2 plants, primarily due to reduced cell elongation (Sheahan et al., 2004). Another study showed that alcohol- inducible expression of GFP-mTalin in root hairs caused severe defects in actin organisation, resulting in either the termination of root hair growth and / or changes in cell shape (Ketelaar et al., 2004). No such observation was recorded in the root cells in particular.

Cytoplasmic aggregation, the rapid translocation of cytoplasm and sub cellular components to the site of pathogen penetration, is one of the earliest reactions of plant cells against attack by microorganisms. During interaction between Arabidopsis and oomycete pathogens GFP-TUA6 and GFP-hTalin were studied. In all interactions, actin microfilaments were actively rearranged and formed large bundles in cytoplasmic strands focused on the penetration site. GFP probes also demonstrated aggregation of ER membrane and accumulation of Golgi bodies at the infection site, suggesting that production and secretion of plant materials were activated around the penetration site. Microtubules did not become focused on the penetration site. No difference was evident between the responses of epidermal cells in the non-host, incompatible and compatible interactions. This result indicates that the induction of cytoplasmic aggregation in Arabidopsis epidermal cells was neither suppressed by the virulent strain of pathogen, nor effective in stopping infection (Takemoto et al., 2003).

Studies such as this demonstrate the suitability of GFP-transformed Arabidopsis for my project to study the effects of nematode parasitism on the cytoskeleton.

2.1.7 Nematodes and the plant cytoskeleton

While infecting a host plant, a plant parasitic nematode can induce changes in the host cell anatomy and physiology, and affect cytoskeleton structure, in order to enable parasitism. Several studies have shown that changes occur in the structure of plant cells in roots infected by nematodes. Due to the fact that most of these studies were conducted using the nematode Meloidogyne incognita, most of the details now known are from host-parasite reaction involving that nematode species.

Large multinucleate “giant cells” were induced in tissues of plant roots infected by Meloidogyne (Jones 1981a, b; De Almeida Engler et al., 1999; 2004). The expression of tubulin and actin in giant cells was up-regulated to allow the assembly of a new cytoskeleton in expanding feeding cells. However, highly-organised microtubules and actin microfilaments were not observed and were partially depolymerised throughout the feeding site development. A disturbed cytoskeleton was still visible in giant cells and a functional mitotic apparatus was present that contained spindles and arrested phragmoplasts but no preprophase bands (De Almeida Engler et al., 2004).

Drugs affecting microtubules (phalloidin, colchicine and latrunculin), and actin microfilaments (taxol and cytochalasin), caused abnormal cellular differentiation in plants. Characteristics of affected cells included changes in size and shape of cells, wall thickenings and polyploidy. These resemble symptoms of Meloidogyne infection on host root cells.

Colchicine-treated protophloem sieve elements (PSEs) have been reported to differentiate into abnormal cell types (Eleftheriou, 1993). In this study, upon colchicine treatment, the PSEs stopped elongating and increased in diameter and showed abnormal ultrastructural features. Microtubules disorganised and disappeared and cell divisions were blocked. Cells became polyploid and nuclei developed multiple lobes and deep invaginations. Crystalline material accumulated in the cytoplasm. Walls developed unusual thickenings. Root growth declined and vascular elements differentiated close to the root apex (Eleftheriou, 1993). Short, swollen and abnormal PSEs have also been observed in other plant species when cell division is disrupted using chemicals (Apostolakos et al., 1991; Galatis, 1991). When mitosis

was blocked using hydroxyurea and oryzalin in nematode-infected cells, gall development was arrested. A contiguous semicircular structure was formed by the nuclei of giant cells (Wiggers et al., 2002; De Almeida Engler et al., 2004).

Chemical stabilization of the microtubular cytoskeleton with taxol blocked feeding site development in Arabidopsis by Meloidogyne incognita. But when actin or the microtubule cytoskeleton was depolymerised by cytochalasin D or oryzalin, nematodes could complete their life cycle, suggesting that the cytoskeletal rearrangements and depolymerisation induced may be essential for a successful feeding process (De Almeida Engler et al., 2004).

Cell death and nuclear movements were inhibited by the actin-targeted drug cytochalasin E, suggesting that microfilaments are required for the hypersensitive response (Škalamera and Heath, 1998). Meloidogyne-resistant varieties of coffee showed hypersensitive reactions including cell senescence and death as part of the host defense response (Anthony et al., 2005). Cytochalasin also affects cytoplasmic streaming in plants (Collings et al., 1995).

The cytoskeleton plays an important role in cell signalling in plants (Volkmann and Baluška, 1999; Heath, 1997; Collings and Allen, 2000). Extensive signalling between the nematode and the plant host enables parasitism (Bird, 2004). When plant parasitic nematodes such as Meloidogyne spp. and Pratylenchus spp. infect the root epidermis, the cytoskeleton might play a pivotal role in relaying the information of the invasion to the rest of the plant.

Similarities between galls and root nodules have been reinforced by the identification of several genes that are common to both nematode parasite and symbiotic bacteria. A host reaction similar to nodule formation was observed in Meloidogyne-infected plant roots resulting in the characteristic galls, enclosing the mature females (Lohar et al, 2004; Hirsch et al, 2002; McCarter et al., 2003; Koltai et al., 2001; Weerasinghe et al., 2005; Bird 1996).

Host microtubules are affected during infection by the nematode shown by the continuous activity of beta tubulin 1 (TUB1), which occurred within giant cells

produced by Meloidogyne incognita in A. thaliana. This was consistent with the activity of a TUB-1 promoter 28 d post infection, which is about the time of first egg production by the nematode (Green et al., 2002). Extensive rearrangement of the cytoskeleton in response to infection of Arabidopsis by Meloidogyne was also observed (De Almeida Engler et al., 2004).

Increased GFP fluorescence was induced in and around feeding sites by Meloidogyne (Hallman et al., 2001; Urwin et al., 1997; De Almeida Engler et al., 2004).

The above studies show that when a host plant is infected by Meloidogyne the host undergoes many modifications. As most normal functions of the cell involve the cytoskeleton, nematode infection would certainly have a great impact on the cytoskeleton. Moreover, wounds caused by the breaching of epidermal cells by plant parasitic nematodes like Meloidogyne and Pratylenchus would also involve the cytoskeleton. Therefore it is important to study the effects of nematode infection on plant roots to gain a better understanding of the infection process. The contrasting modes of parasitism (obligatory endoparasitism of Meloidogyne and migratory parasitism of Pratylenchus) will shed light on the infection processes and host responses from two different standpoints.

Literature was difficult to find on cytoskeletal studies using Pratylenchus, and during my search no literature could be found of any previous study on the subject. This will be the first study of the cytoskeletal aspects of early stages of parasitism of Pratylenchus on Arabidopsis.

2.2 Materials and Methods

2.2.1 Culture of Meloidogyne javanica inoculum

Tomato, S. lycopersicon cv. Tiny Tim, seeds were obtained from Dr Jennifer Cobon (DPI, Indooroopilly, Queensland). These seeds were surface sterilised by treating with 70% v/v ethanol (15 min) followed by 2.5% NaOCl v/v with Tween-20 (one drop per 100 mL for 20-30 min). Liquid and seeds were then poured through a sterile

sieve and passed through four washes of sterile distilled water. Using a sterile forceps, tomato seeds were placed onto the surface of agar in 9 cm diameter deep Petri dishes with at least 6 mm solid growth medium (quarter-strength Murashige and Skoog medium supplemented by 0.5% sucrose and solidified with 0.6% Phytagel at pH 6.4; Hutangura et al. (1998). Fourteen seeds were distributed randomly in each Petri dish, which were sealed with Nescofilm and stored at 4o C until needed. When necessary, they were placed in growth cabinets with a 16 h light / 8 h dark cycle at a constant temperature of 27° C. After 10-14 d, each Petri dish was inoculated with approximately 100 µL of sterile water containing 300 Meloidogyne second stage larvae (J2) and sealed and incubated in the same cabinet. (Initial inoculum of Meloidogyne for the purpose of the study was obtained from Dr. Zhaohui Wang, Murdoch University, Perth, WA). Galls on tomato were observed from the third day after inoculation. After 4-6 weeks, egg masses could be observed. Sterile egg masses were removed under aseptic conditions and placed in a sterile water droplet in a small (3-5 cm diameter) sterile Petri dish. Approximately 4-5 egg masses were placed in each droplet. The Petri dish was sealed with Nescofilm and incubated in the previously mentioned growth cabinet. J2 hatched out in 2 d, and the liquid containing these J2 was used for inoculating aseptic tomato seedlings to maintain the inoculum or to inoculate Arabidopsis seedlings for the experiment. Fresh sterile water was added and more J2 were observed to hatch out in a further 2 d (Hutangura et al., 1998; 1999).

2.2.2 Pratylenchus thornei inoculum

P. thornei cultures maintained on carrots were obtained from Dr Jennifer Cobon (DPI Queensland) and Dr Sharyn Taylor (SARDI, South Australia) and used for the study. Aseptic carrot pieces containing Pratylenchus were placed on a sterile glass plate in a laminar flow cabinet and small pieces were cut from it. The cut pieces of carrot were incubated for 24 h in sterile water in a petridish at 21° C to extract the larvae. The liquid containing the Pratylenchus larvae was used to inoculate Arabidopsis seedlings for the experiment. The original container was resealed aseptically for extracting nematodes when necessary for subsequent experiments.

2.2.3 Culture of Arabidopsis thaliana ecotype Columbia as host

Arabidopsis seeds were surface sterilised by treating with 70% v/v ethanol (2 min) followed by 2.5% NaOCl v/v with a drop of Tween-20 (one drop per 100 mL, 2 min). Seeds were washed in four changes of sterile distilled water. A row of 10 single seeds was then planted on to the agar surface (Hutangura et al., 1998) in a 9 cm Petri dish. Seeded plates were stored at 4° C in a cool room for a minimum of 48 h to synchronise germination. As required, plates were then placed in a growth cabinet with a 16 h light / 8 h dark cycle at a constant 27° C for subsequent infection studies.

2.2.4 Immunolabelling for confocal microscopy of whole roots

The following procedures used by Gardiner et al. (2003) and Collings and Wasteneys (2005) were followed to immunolabel the cytoskeleton of Arabidopsis roots. Whole Arabidopsis seedlings, whether infected or uninfected, were fixed in PME buffer [50 + mM PIPES pH 7.2 (K ), 2 mM EGTA, 2 mM MgSO4 and 0.1% (v/v) Triton X-100] containing 200 mM PMSF in ethanol, 40 mM MBS in DMSO, 4% (v/v) formaldehyde and 1% (v/v) glutaraldehyde (Sigma) for 40 min and washed in PME 3 times for 5 min each. Then they were extracted (60 min) in PME containing 1% (v/v) Triton X-100 and again washed in PME buffer. After that their cell walls were digested (20 min) with an enzyme mixture (1% [w/v] cellulase Y6 and 0.1% [w/v] pectolyase Y23 [ICN, Seven Hills, NSW, Australia]) dissolved in PME buffer containing 1% (w/v) BSA (Fraction V, Sigma, Sydney, NSW, Australia) and 0.4% w/v) Mannitol. They were then washed (2 X 10 min) in PME buffer. Seedlings were then permeabilised with 100% (v/v) methanol at -200C and then rehydrated in PBS

(131 mM NaCl, 5.1 mM NaH2PO4, and 1.56 mM KH2PO4, pH 7.2) for 10 min. After washing (3 X 10 min) each in PBS, free aldehyde groups were reduced with freshly bubbling sodium borohydride at 5 mg/mL in PBS (20 min). Seedlings were washed in PBS and placed on to 3 or 5 cm diameter Petri dishes lined with a layer of Nescofilm. 50µL of incubation buffer (PBS containing 1% BSA and 50 mM glycine) was added (30 min) to block non-specific antibody-binding sites. The Petri dish was gently tilted and the liquid drawn off using a tissue paper. Material was then washed in PBS-glycine 3 X 20 min and incubated in primary antibody (polyclonal anti-actin

1/200 from Chris Staiger (Purdue University, West Lafayette, Indiana, USA) and monoclonal anti-tubulin 1/1000 from Sigma clone B512 diluted 1/200 and 1/1000 respectively in incubation buffer) (2 h) at room temperature. After removing primary antibody, seedlings were washed in PBS containing 50 mM glycine and then incubated for 3 h in secondary antibody (sheep-antirabbit FITC (Silenus Boronia, Victoria) and goat-anti mouse Cy-5 (Jackson, West Grove, PA, USA) diluted 1/100 and 1/200 respectively in incubation buffer). They were washed in PBS-Glycine (3 X 10 min) and incubated in 1 µg / mL DAPI (4’, 6-diamidino-2-phenylindole hydrochloride) Sigma (10 min). After this they were lifted onto a drop of Citifluor (Citifluor, London, U.K.) on a microscopic slide after placing a drop of nail polish at each of the four corners of the slide to prevent the seedlings from being crushed. A cover glass was placed carefully on top and sealed using nail polish. Samples were viewed with 60X or 40X oil immersion lenses on a Leica SP2 confocal microscope.

2.2.5 Embedding and sectioning of whole roots in BMM

The method described by Baskin et al. (1992) was followed for this procedure, with slight modifications. Seedlings were fixed as described in Section 2.2.4. A portion of the root containing the gall was excised and passed successively through 10, 25, 50, 75, 95 and 100% (v/v) solution of ethanol in distilled water (30 min each). Two more changes of 100% ethanol were given and the samples were left overnight in the final change of 100% ethanol after adding 1µL / mL of 8% Fast Green stain to make the roots more visible for later processing. Dehydration steps were done at 4o C. Tissue was then infiltrated with 2:1, 1:1 and 1:2 ethanol: resin mixture (BMM resin: 4:1 (v/v) butyl methacrylate to methyl methacrylate, to which was added 0.5% (w/v) benzoin ethyl ether) (2 h each). After passing the tissue through two changes of 100% BMM resin (2 h each), it was left in 100% resin for at least 24 h. Infiltration steps were done at 4° C or - 20°C. Tissue was then placed on aluminium foil trays and covered with fresh resin. The trays were then placed in the polymerisation chamber, flushed with N2 gas, UV lights turned on and the sample left to polymerise overnight. Parts of the polymerised block containing root samples were cut into suitable sized cubes using a saw and were stuck to a plastic stub using a drop of araldite (Selleys Araldite Ultra Clear®, Selleys Pty Ltd, Padstow NSW, Australia). About 30 min later (to allow for drying of araldite), dry sections of varying

thicknesses (10-50 µm) were cut using a glass knife on an ultra microtome. Sections were floated on to drops of water on 12-welled glass slides coated with PEI (polyethylenimine) and air dried at room temperature on the laboratory bench.

2.2.6 Immunolabelling BMM sections for confocal microscopy

Selected slides were immersed in 100% acetone in a Coplin jar for 10 min to remove embedding resin, and washed in PBS (3 X 5 min), sodium borohydride (5 mg/mL in PBS) (10 min), and washed in PBS (3 X 5 min). Slides were then taken out and 20 µL of incubation buffer (PBS containing 1% [w/v] BSA and 50 mM glycine) was put on each slide well containing sections for 5 min. The slides were then tilted on to a tissue paper to draw off the incubation buffer. To each slide well 20 µL of primary antibody (Section 2.2.4) was added and incubated for 2 h at room temperature, washed in PBS-glycine (3 X 5 min), then treated with secondary antibody (2 h). After washing in PBS-glycine the sections were stained for 10 min with DAPI, mounted in Citifluor and sealed with nail polish and examined with a 40X oil immersion lens on the Leica SP2 Confocal Microscope.

2.2.7 Visualisation of the actin cytoskeleton in living root cells of Arabidopsis thaliana using GFP-hTalin fusion protein

To study changes in the actin cytoskeleton due to infection by Meloidogyne in live plants, a transgenic A. thaliana line expressing a Green Fluorescent Protein (GFP)- tagged h-Talin reporter protein was used. The specific line of transgenic Arabidopsis plants expressing GFP-h Talin was obtained from Daigo Takemoto (Plant Cell Biology Group, RSBS, ANU) to study actin microfilaments. Plants were grown as described in Section 2.2.3. For observation they were mounted in water on a glass slide and covered with a cover glass. GFP fluorescence in these plants was then observed concurrently with transmission light images with a 63X water immersion lens using a Leica SP2 confocal microscope.

2.3 Results

The cytoskeleton was initially characterised in A. thaliana in uninfected roots and then in roots following infection with M. javanica and P. thornei. The following methods were used to compare the cytoskeleton and its changes.

1) Whole root immunolabelling (Collings and Wasteneys, 2005), 2) Immunolabelling of BMM sections (Baskin et al., 1992), and, 3) Imaging of living tissue using GFP-hTalin (Takemoto et al., 2003)

In images showing microtubules and actin filaments in a single image (as an overlay), microtubules are false-coloured red and actin filaments green. Antibody controls in which primary antibodies were removed but secondary antibodies were used alone, demonstrated that the actin and tubulin labelling patterns found with the primary antibodies were specific (data not shown). This is the first published study to use whole root dual immunolabelling to study nematode infection on plant roots.

2.3.1 Uninfected Arabidopsis thaliana roots

2.3.1.1 Root cap and meristem immunolabelled for actin and microtubules

Of the 120 uninfected plants processed for immunolabelling, 102 (85%) showed successful labelling of the cytoskeleton in root cells. Labelling was apparent in the epidermal cells of the meristem and elongation zones, and in some cases, even root hairs were labelled (Figures 2.03 to 2.05). In general however, it was difficult to label cells in the mature regions of the root. The likely reason for this was the difficulty in getting antibodies to penetrate the cell wall once cells had ceased elongating, and secondary wall thickening had commenced as cells in this region ceased to elongate. Studies of actin filaments were difficult because these are much harder to fix than microtubules. It was also found that adequate fixing, especially of actin filaments, was not possible unless samples were immersed in the fixative solution within 2 to 3 min of making the solution. Where proper fixation of actin did not occur, labelling of actin appeared only as a diffuse fluorescence. When fixation was adequate; however, actin bundles were observed in all cells of uninfected

Arabidopsis roots. Overcrowding of the fixative solution also resulted in poor fixation. For good fixation the maximum number of samples per mL fixative solution was found to be six or less.

The Arabidopsis root tip is composed of many cell layers. Starting from the outermost layer, these are the root cap cells that sheathe the growing tip of the root, then the epidermis, cortex, endodermis, and the innermost section, the stele or vascular tissue (Dolan et al., 1993; Bowman, 1994). In the elongating region of the root further behind the root tip, the root cap cells may slough off leaving the epidermal cells exposed. Root cap cells were readily labelled and more fluorescent. The root tip or meristem consisted of actively dividing and growing meristematic cells. In the epidermis of the root tip of uninfected Arabidopsis, numerous dividing cells were observed, in various stages of division (Figures 2.03A, 2.03B). Oblong plant cells were arranged regularly in roots with their long axis parallel to the long axis of the root. The root epidermis consisted of two types of cells: root hair cells (trichoblasts) and non-root hair cells (atrichoblasts). Trichoblasts are located over the anticlinal wall between two cortical cells. One, or in some cases two, atrichoblast files were located in between two files of trichoblasts. Root hairs develop from the trichoblast cells only after the cells have ceased to elongate. The alternating fluorescent and non-fluorescent trichoblast and atrichoblast cell files could clearly be seen (Figures 2.10 and 2.11).

Figure 2.03A An Arabidopsis thaliana root triple labelled for actin, microtubules and DNA shows the different stages of cell division in an uninfected root meristem.

Confocal optical sections show actin (green), microtubules (red) and DNA (blue) at 4 µm (A) and 18 µm (B) from the surface of the root. The different stages of the cell cycle, defined by different microtubule arrays, were visible: ppb, pre-prophase band; sp, mitotic spindle; ph, phragmoplast; Tmt, transversely-arranged interphase microtubules. Other abbreviations: I, cell in interphase; P, cell in prophase; M, cell in metaphase; T, two cells in telophase with the phragmoplast laying down the cell wall between two daughter nuclei (the two adjacent cells marked T are dividing perpendicular to each other); a, actin; c, chromosomes arranged in the metaphase plate; D, a cell in transition between prophase and metaphase showing both preprophase band and a newly forming spindle. A Cells at interphase showed transverse microtubules (Tmt). In cells in the stage just before prophase the microtubules formed a pre-prophase band (ppb) here seen as a ring around the nucleus (N). In interphase cells, actin microfilaments were mainly endoplasmic (a). B Several cells in various stages of division (interphase, prophase, an intermediate stage between prophase and metaphase, metaphase and telophase) emphasised the different roles of actin microfilaments and microtubules. A cell in transition between prophase and metaphase (D) shows the depolymerisation of the microtubules in the preprophase band and their rearrangement in the formation of the spindle. Chromosomes are arranged in the metaphase plate in a cell in metaphase (M); Actin, seen along with microtubules, in the phragmoplast (ph) in two adjacent cells in telophase (T) which are dividing at angles perpendicular to each other. This process occurred in an older plant cell, which has resulted in one cell file then giving rise to two cell files (cf). Bar = 20 µm.

Figure 2.03B An Arabidopsis thaliana root triple labelled for actin, microtubules and DNA shows the different stages of cell division in an uninfected root meristem.

In interphase cells, actin filaments were not observed in the cortical region of the cell (Figure 2.03); they were endoplasmic, mainly radiating through the cytoplasm. Some actin filaments and bundles were also arranged parallel to the longitudinal axis of the root.

Figure 2.04A An uninfected Arabidopsis thaliana root shows actin organisation in the post-mitotic root tip.

Confocal immunolabelling of actin (A) and microtubules (B), shown as maximum projections of 38 images taken at 1 µm intervals. A Actin bundles (ab) in root tip were mainly organised longitudinal. B Microtubule organisation in the outer epidermal layer was transverse (Tmt) whereas cells in the inner cortical layer had oblique or random microtubules. Bar = 30 m.

Figure 2.04B An uninfected Arabidopsis thaliana root shows microtubule organisation in the post-mitotic root tip.

Actin bundles and filaments were observed throughout the cytoplasm, sometimes forming a network around the nucleus (Figure 2.04A).

In the early stages of cell division, actin filaments were very rarely co-localised with endoplasmic microtubules, or in the cortical region lining the cell wall. In later stages of division, actin filaments co-localised with microtubules in the phragmoplast, which lays down the new cell wall that divides the daughter cells.

Microtubules were observed in all cells of Arabidopsis roots (Figures 2.03 to 2.05), except in those cells that were not properly fixed. Details of cytoskeletal involvement in the different stages of the cell cycle are given in Chapter 1 of this thesis.

Changes in microtubule organisation were observed through the cell cycle. In interphase cells of the root meristem region, cortical microtubules were arranged transverse to the longitudinal axis of the root (Figure 2.03). Further into the cell, microtubules were randomly arranged in the cytoplasm, sometimes even longitudinal, parallel to the axis of the root (Figure 2.04). Root expansion in the meristem, resulting in an increase in root diameter, was apparent from occasional longitudinal cell divisions that give rise to two cell files (Figure 2.04B). In some cases, adjacent cells in telophase divided at right angles to each other (Figure 2.03B). Behind the meristem there was little or no increase in root width. Sometimes it appeared that actin bundles in two adjacent cells mirrored each other, giving the impression of a continuous strand between the two adjacent cells (Figure 2.04A).

2.3.1.2 Immunolabelling the differentiating region of whole roots for actin and microtubules

In the cells of this region, actin filament distribution was mostly endoplasmic (Figure 2.05A) with some bundles running obliquely and some almost parallel to the longitudinal axis of the root. Many interconnections were observed between these bundles. Cells of the stele were long and narrow, so the actin in these cells, when observed, appeared to be longitudinal.

Figure 2.05 The organisation of actin and microtubules in differentiating cells of an uninfected A. thaliana root.

Confocal immunolabelling of actin (A) and microtubules (B), shown as maximum projections of 129 images taken at 1 µm intervals from the root surface. The long axis of each cell is oriented parallel to the long axis of the root. A Actin bundles (ab) in the differentiating region of the root were mainly longitudinal or oblique. B Microtubules were arranged in an oblique direction (Omt). Microtubule organisation was clearly visible in a root hair (rh). Bar = 30 m.

Cell divisions did not occur in the epidermis of the differentiating region in the root. As a result, microtubular structures observed normally in the meristematic region were not observed in cells in the differentiating region. In the cells in this region, microtubules were cortical and arranged obliquely in relation to the longitudinal axis of the root (Figure 2.05B).

2.3.1.3 Immunolabelling the cytoskeleton in BMM sections of roots

In order to compare results obtained through whole root immunolabelling, I attempted to optimise the method described by Baskin et al. (1992) for embedding, sectioning and immunolabelling nematode-infected and uninfected Arabidopsis roots in BMM (butyl methyl methacrylate) resin. Only root sections from the differentiating region of the root were embedded, as these would form the basis for comparisons with galls in Arabidopsis roots infected with Meloidogyne.

Figure 2.06A Immunolabelling of a BMM section of an uninfected Arabidopsis thaliana root.

Confocal immunolabelling of actin (Figure 2.06A) and microtubules (Figure 2.06B), shown as maximum projections of 20 images taken at 1 µm intervals. A Actin bundle (ab) in the cells of the vascular bundle (vb) arranged longitudinally in the root; in one cell actin was observed in a phragmoplast (ph). This division was likely in the pericycle which remains mitotically active even after divisions cease in the mature root regions. B Microtubules were generally oblique (Omt) and were also present in the phragmoplast (ph). Bar = 20 µm.

Figure 2.06B Immunolabelling of a BMM section of an uninfected Arabidopsis thaliana root.

More actin labelling was visible in the stele than in other cell layers (Figure 2.06A). Large bundles of actin filaments were oriented parallel to the longitudinal axis of the root. One section showed a very rare cell division in the vascular region, probably in the pericycle in which actin and microtubules were visible in a phragmoplast. In longitudinal sections of the differentiating region of the Arabidopsis root, microtubules were cortical and arranged obliquely in relation to the longitudinal axis of the root (Figure 2.06B) indicating that elongation had ceased.

2.3.1.4 The actin cytoskeleton in living Arabidopsis thaliana roots

I investigated actin filament organisation in living, uninfected Arabidopsis roots using a line of Arabidopsis plants transformed to stably express a fusion protein of GFP and the actin-binding domain of human talin (GFP-hTalin). Seeds were kindly provided by Daigo Takemoto and David Jones (RSBS, ANU) (Takemoto et al., 2003). The expression of GFP-hTalin was predominantly in the meristem and stele, areas that I was interested in studying. Due to the minimal processing involved, using GFP-hTalin plants was the easiest and least time consuming way of studying the actin cytoskeleton. The living plants were observed by confocal microscopy under blue light so that fluorescence from GFP would showed the localisation of talin, and thus actin.

A large percentage (about 90%) of the germinated GFP-hTalin plants showed fluorescence. [A possible explanation why 10% of the plants did not show fluorescence is that some sort of gene silencing occurred in the talin lines so that the transcription and/or translation of the GFP-hTalin were prevented (Dr. David Collings, Personal Communication)]. Fluorescence was mainly found in the root cap, but behind the root cap, the quiescent centre did not show fluorescence. Beyond this region strong fluorescence was observed in the stele (Figure 2.07). In the meristem, several cell layers forming a ring around the central cylinder were more fluorescent than the surrounding cortical or central cell files.

These tissues undergo development and differentiation and subsequently form the vascular bundle. Actin was observed in the elongating and differentiating region of the Arabidopsis root. Nuclei in the epidermal cells were surrounded by actin, which radiated out to the plasma membrane. Although actin was mainly longitudinal to the axis of the root, interconnections were observed between adjacent bundles. This sometimes gave the impression of a transverse arrangement (Figure 2.08). Actin filaments also extended into the root hair where they formed a network (Figure 2.08, arh).

Figure 2.07 A GFP-hTalin-transformed Arabidopsis thaliana seedling showing actin in the developing vasculature of the root tip.

GFP-hTalin showed the strongest actin labelling in the developing vascular tissue (stele) and root cap, although weaker labelling was also present in epidermal and cortical cells. The image is a maximum projection of confocal optical sections taken from 20 µm to 24 µm from the surface at 1 µm intervals. Bar = 30 µm.

Figure 2.08 GFP-hTalin labelling revealed an extensive actin cytoskeleton in the epidermis of the root differentiation zone.

GFP-hTalin revealed actin organisation in the differentiating region of the root. Nuclei are surrounded by an actin network (asn) and actin (ab) radiated from nuclei to the plasma membrane. Actin was also observed in root hairs (arh). Bar = 10 µm.

It was my intention to compare microtubules in nematode-infected and uninfected plants that stably expressed GFP-MBD (Arabidopsis expressing GFP fused with the microtubule-binding domain of mouse MAP4) plants (Marc et al., 1998). However, unfortunately these plants lacked detectable fluorescence in the stele, and therefore the study could not be conducted.

2.3.2 Arabidopsis thaliana infected by Meloidogyne javanica

As the individual nematodes, whether Meloidogyne or Pratylenchus, were found to infect the plants at different times after inoculation, it was easier to divide the infection process into three processes and time periods: (i) Nematode entry (ii) Nematode migration within the root (iii) Nematode feeding sites

2.3.2.1 Immunolabelling whole roots to observe actin filaments and microtubules during Meloidogyne javanica entry

Aseptically-cultured M. javanica were used to inoculate four-day-old A. thaliana seedlings. Inoculated plant roots were observed daily. However, as individual nematodes entered the root at different times after inoculation, gall formation times also varied. Indeed, nematodes remained swimming around in the root zone even a week after inoculation. From the first observation 3 h after inoculation, highly fluorescent material that labelled with actin antibodies was extruded from around the wound site (Figure 2.09 A-C). This fluorescence could be distinguished from autofluorescence by the distinctive colour. Thus, it was always easy to find a wound on the surface of the root by the presence of dense green fluorescence of actin above the surface of the wounded cell. Autofluorescence seemed to be represented by a slightly more yellowish green colour.

Microtubules were disrupted or absent from the wound (Figure 2.09 F-H). Nematodes themselves were not observed in the surface wounds in most cases. However, in one instance a nematode was found with its head inside root tissue although the body of the nematode was destroyed during immunolabelling (Figure 2.09 I-J).

Figure 2.09 Immunolabelling of a Meloidogyne-infected Arabidopsis root demonstrated cytoskeletal disruption during nematode entry.

An Arabidopsis root infected with M. javanica was concurrently immunolabelled for actin and microtubules 24 h after inoculation. Confocal microscope images showed the organisation of actin (A- E) and microtubules (F-J). A, F Maximum projections of 16 images at 1 µm intervals. B-E, G-J selected optical sections at 4 µm intervals from the root surface. An arrow (E) points to a nematode (n) and a cell dislodged near nematode entry region; arrowheads show fluorescent labelling on the surface of root and in the wound area that was positive for actin and tubulin antibodies. Bar = 100µm.

Figure 2.10 Immunolabelling of a Meloidogyne-infected Arabidopsis root demonstrated cytoskeletal disruption adjacent to the root tip during nematode entry.

An Arabidopsis thaliana root infected with Meloidogyne javanica concurrently labelled for actin and microtubules 24 h after inoculation showed organisation of actin (A-D) and microtubules (E-H) during nematode entry just behind the root tip. I-K Concurrent transmitted light images. A, E Maximum projections of 30 sections taken at 1 µm intervals. B-D, F-H, I-K Selected optical sections at 10 µm intervals from the root surface. Increased actin and microtubule labelling occurred around wounds. Arrowheads point to the wound; single arrows point to the nematode which was destroyed during processing; double headed arrows show alternating files of fluorescent and non-fluorescent cells which are indicators of future root hair and non-root hair cells. Bar = 20 µm.

Figure 2.11 An Arabidopsis thaliana root tip infected with Meloidogyne javanica showed increased cytoskeletal labelling and disruption.

An Arabidopsis root tip infected with Meloidogyne concurrently labelled for actin and microtubules 24 h after inoculation showed actin (A-E) and microtubules (F-J) in the root during the process of nematode entry through the tip. A, F Maximum projections of 50 sections taken at 1 µm intervals. B- E, G-J, K-O Selected optical sections at 10 µm intervals from the root surface. Arrowheads show increased fluorescence in cells surrounding entry wound; short arrows point to the body of the nematode; triple arrow points to alternating fluorescent and non-fluorescent trichoblasts and atrichoblasts. Root hairs (rh) have formed close to the root tip due to slowing of root growth because of nematode infection Bar = 100 µm.

Alternating fluorescent and non-fluorescent trichoblast and atrichoblast cell files were present (Figures 2.10 B-D and F-H), even when tissues were severely disrupted due to nematode invasion (Figure 2.11 A, F).

In the majority of infected roots observed (97%), Meloidogyne juveniles entered the root behind the root tip, mainly in the elongating region, but in 3% of cases entry was through the root tip apex. In root tip apex entries, cell files were disturbed to such an extent that no normal root tip tissue was recognisable and neither the root cap nor the quiescent centre was clearly discernible (Figure 2.11). In the infection site, cells were intensely labelled for actin and microtubules, especially the cells adjacent to the nematode body (Figure 2.11 C, H). In cases where the root tip was infected, the rate of growth appeared to have slowed down considerably so that large numbers of root hairs were found concentrated very close to the root tip because of the compression of the root elongation zone (Figure 2.11 E, O). In all 300 infected roots observed, tissue was more fluorescent near the wound site than the surrounding cells. As mentioned above, autofluorescent cells had a slightly yellowish green fluorescence compared to the bright green labelling of actin.

2.3.2.2 Immunolabelling whole root to observe actin and microtubules during Meloidogyne migration within the root

During the initial migration of the nematode towards the tip of the root, cell shape in the path of the nematode was not affected. Cells were not displaced from their normal arrangement. (Figure 2.12). Nematodes migrated towards the root tip outside the vascular bundle parallel to the long axis of the root (Figure 2.12 E-G).

Figure 2.12 Cytoskeletal changes during the migration of Meloidogyne javanica juveniles within an Arabidopsis thaliana root.

Confocal immunolabelling of an Arabidopsis thaliana root infected with Meloidogyne javanica showed organisation of microtubules during nematode migration. A maximum projection of 12 sections taken at 1 µm intervals. B-D, Three optical sections at 6 µm intervals from the root surface. E-G Concurrent transmitted light images. Arrowheads point to Meloidogyne juveniles migrating within root; arrows point to characteristic wavy microtubules in root cells adjacent to the nematode. Bar = 20 µm.

Microtubules close to the body of the migrating nematode were more strongly labelled than the surrounding cells (Figure 2.12 A-D). The microtubules did not show their normal taut arrangement but were loose and wavy. Elements of nematode morphology including the stylet, oesophagus, and even the crenulations on the cuticular surface were visible (Figure 2.13 C, D).

Figure 2.13 Magnified view of the cytoskeletal changes during the migration of Meloidogyne javanica juveniles within an Arabidopsis thaliana root.

An Arabidopsis thaliana root infected with Meloidogyne javanica immunolabelled for microtubules showed their organisation in root cells adjacent to migrating juveniles. Confocal immunolabelling for microtubules (A, B); transmitted light images (C, D). A Maximum projection of 27 images while C is an optical section at 27 µm from the root surface. B, D Sections at 6 µm from the root surface. Arrows in A and B show wavy microtubules. Arrowheads (C, D) indicate Meloidogyne javanica juveniles. Bar = 30 µm.

Figure 2.14 An Arabidopsis thaliana root infected with Meloidogyne javanica showing microtubule organisation in root tip cells close to the nematode.

A-F Confocal immunolabelling of microtubules G-K Transmitted light images B-F, G-K Individual sections at 1, 5, 9, 13 and 35 µm from the root surface. A maximum projection of 35 sections. Images show the nematode in the root tip turning into the vascular bundle. The arrow points to loose and wavy microtubules in a cell adjacent to the nematode body that was dislodged from its normal position. Bar = 20 µm.

Nematodes, having reached the root tip, turned and entered the differentiating vascular bundle (Figure 2.14). Cortical microtubules in a cell in the infected meristematic region adjacent to the migrating nematode appeared relaxed and wavy instead of being of a more typical tightly stretched, transverse arrangement (Figure 2.14 A). This cell seemed to be displaced, probably due to the mechanical pressure exerted by the turning nematode.

The rate of growth of the root tip was so severely reduced by nematode infection that the xylem elements differentiated immediately behind the root cap (Figure 2.14 K).

Roots were inoculated near the root tip and of the 300 infected samples observed about 5% of roots showed root tip galls. This was due to a large number of nematodes infecting the same area of the root tip simultaneously (Figure 2.15 F-H).

Figure 2.15 Immunolabelling of a root tip gall in Arabidopsis thaliana infected with Meloidogyne javanica showing the organisation of microtubules in the gall tissue.

An Arabidopsis thaliana root tip gall (the root tip was infected with multiple M. javanica juveniles) immunolabelled for microtubules (A-E). A Maximum projection of 47 sections taken at 1 µm intervals; B-E Maximum projections of 11 optical sections at 1-11, 12-23, 24-35 and 36-47 µm from the root surface respectively. F-I Concurrent transmitted light images taken at 11, 23, 35 and 47 µm from the root surface. Transmitted light showed several nematodes in the gall with arrowheads pointing to M. javanica juveniles. Cells in the gall (G) had increased labelling of microtubules compared to neighbouring cells. Many irregularly-shaped, enlarged cells were observed in the gall tissue with these possibly being the giant cells (gc) induced by the nematodes. Most cells in the gall seemed to have lost their polarity of growth (clp). The cells with lost polarity were positioned with their axes pointed in various different directions. Bar = 30 µm.

In these cases, extensive proliferation of cells was observed, resulting in single or multiple galls. The cells in these galls were smaller in size and larger in number and more of a spherical shape than uninfected plant cells (Figure 2.15 A, B). In all cases the cells adjacent to the nematode, especially those in the gall tissue showed increased fluorescence (Figure 2.15 A-E).

2.3.2.3 Immunolabelling whole roots to observe actin filaments and microtubules in Meloidogyne javanica feeding sites

Two days after inoculation, small bulges began to form on the roots indicating feeding site formation. These galls were in the root differentiation zone. Examination of galls revealed increased labelling of the cytoskeleton in cells in the feeding site, especially in the vascular region (Figure 2.16). Though the galls usually formed in the elongating region, by the time of observation, cells would already be differentiated in the infected region due to continuous root growth.

Cells in the observed gall appeared to be abnormally spherical or elongated compared to the shape of a normal plant cell. In these cells comprising the feeding site in the differentiated region, microtubules were mostly transverse, rather than the usual oblique arrangement of microtubules in uninfected cells of the differentiating region (Figure 2.16 A-F). Cells in the gall had lost their polarity, and were aligned in various directions instead of the normal direction parallel to the long axis of the root. While some gall cells seemed to be positioned with their long axes perpendicular to the long axis of the root, most other cells were positioned at an angle (Figure 2.16 B- D). Numerous cell divisions with intensely labelled spindles occurred in the root all around the body of the nematode, particularly around its posterior (Figure 2.16 F). Plant cells near the head of the nematode were larger. In some cells, unknown microtubular structures, which were thick and semicircular, were observed. These could be incomplete phragmoplasts (Figure 2.16 E). In transmitted light images, optical properties of the cell walls in the infected region appeared to be different from cell walls in neighbouring cells. Cell walls of galls observed in transmitted light were darker and appeared to be thicker than those of the surrounding cells (Figure 2.16 H, I).

Figure 2.16 Microtubule labelling in a two-day-old gall in an Arabidopsis thaliana root infected with Meloidogyne javanica.

Confocal immunolabelling showed microtubules (A-F) in giant cells induced by Meloidogyne javanica in Arabidopsis. These images were collected as optical sections at 1 µm intervals. A Maximum projection of 50 sections. B-F Maximum projections of 10 sections each, taken at 1-10, 11- 20, 21-30, 31-40 and 41-50 µm respectively from the root surface. G-K Concurrent transmitted light images taken at 10, 20, 30, 40 and 50 µm from the root surface. In transmitted light, the optical properties of infected cells were different to neighbouring cells especially when they are slightly out- of-focus (cw), possibly because of thicker cell walls. Arrows (A, E) indicate abnormal or incomplete phragmoplasts; sp, numerous spindles showing cell divisions adjacent to the nematode body; gc, giant cell; Tmt, transversely oriented microtubules; clp, cells which have lost their growth polarity; n, nematode within the vascular bundle of the root; sn, a nematode lying on the surface of the root; x, xylem. Bar = 50 µm.

Four days after inoculation, galls had increased in size. The cells in these galls were irregularly spherical in shape and microtubules were arranged transversely along the cell wall (Figure 2.17). Differences in optical properties of infected cells were more pronounced in transmitted light images (Figure 2.17 G, H).

Cell walls of the infected area seemed darker and thicker than surrounding cells. Many spindles (Figure 2.17 E) and an incomplete phragmoplast (Figure 2.17 C) were observed in cells of the gall. Close to the head of the nematode, a broken xylem vessel was observed with transmitted light (Figure 2.17 H, I). Massive disruption of xylem vessels was observed in infected roots, especially in galls, and newly differentiating normal xylem vessels were also observed in these areas of disruption (Figure 2.18).

In galls observed 14 d after inoculation, cellular structure seemed to have degenerated. Actin labelling was mainly observed towards the cell walls (Figure 2.19 A-J).

Figure 2.17 A four-day-old gall in an Arabidopsis thaliana root infected with Meloidogyne javanica immunolabelled for microtubules.

Confocal immunolabelling showed the arrangement of microtubules (A-E) in a four-day old gall. A Maximum projection of 97 optical sections; B-E Single optical sections at 1, 26, 52 and 97 µm from the root surface. F-I Concurrent transmitted light images. Tmt, transverse microtubules in a giant cell; gc, giant cell; sp, spindles in dividing cells of the gall; arrowhead (C) points to an abnormal phragmoplast; n, two nematodes (H, I) within the vascular bundle; x, disrupted xylem vessels; cw, cells in gall tissue showing different optical properties appearing darker compared to neighbouring cells in the root possibly due to thicker cell walls; sp, spindles in dividing cells of the gall in the differentiating region of the root; f, increased microtubule labelling within the gall. Bar = 50 µm.

Figure 2.18 Disruption of xylem in an Arabidopsis thaliana root infected with Meloidogyne javanica.

Transmitted light image 39 µm below the surface of an Arabidopsis thaliana root infected with Meloidogyne javanica. xd, disrupted and malformed xylem vessels in a gall; x, normal xylem differentiating along the length of the root. Bar = 50 µm.

Figure 2.19 A 14-day-old gall induced by Meloidogyne javanica in Arabidopsis thaliana shows deterioration of the cytoskeleton in root tissue.

Concurrent immunolabelling of actin (A-E) and microtubules (F-J) showed their deteriorating structure (with the cytoskeletal structures being less distinct) in a 14-day old gall. A, F Maximum projections of 40 sections taken at intervals of 1 µm from the root surface. B-E, G-J Maximum projections of confocal optical sections taken 1-10, 11-20, 21-30 and 31-40 µm from the root surface respectively. K-N Transmitted light images taken at 10, 20, 30 and 40 µm from the root surface. clp, cells which have lost polarity; a, actin in cells of the gall; Tmt, transverse microtubules in giant cells; Wmt, wavy microtubules observed in some cells. Bar = 50 µm.

Microtubules retained their transverse orientation but the individual bundles of microtubules seemed to be shorter and more disorganised than in galls observed four days after inoculation (Figure 2.19 F-J). Cells of the gall showed varying polarity (Figure 2.19 F, G). Wavy microtubules were also observed (Figure 2.19 H). Galls more than 14 d old were too large for microscopy and good confocal or transmitted light images could not be obtained.

2.3.2.4 Immunolabelling BMM sections of Arabidopsis thaliana root infected by Meloidogyne javanica

Cross sections of galls 14 d after inoculation showed the body of the nematode surrounded by plant cells (Figure 2.20). Infected cells appeared to have co-localised cortical actin and microtubules. Neighbouring uninfected cells showed transverse microtubules while endoplasmic actin filaments and bundles were parallel to the longitudinal axis of the root.

Figure 2.20 Immunolabelling of BMM section of an Arabidopsis thaliana infected by Meloidogyne javanica showing the organisation of actin and microtubules in a root gall.

Confocal optical sections show the arrangement of actin (green) and microtubules (red). Actin and microtubules appear co localised in some areas (yellow). n, nematode body; a, actin; mt, microtubules; col, co-localised actin and microtubules. Bar = 20 µm.

At higher magnification, an irregularly-shaped cell in the infected region showed transverse microtubules and long endoplasmic actin filaments perpendicular to the microtubules. Actin bundles were interconnected with one another (Figure 2.21). In all cases, more fluorescence was observed in cells within and closely adjacent to nematode feeding sites.

Figure 2.21 A BMM section of a Meloidogyne javanica-induced Arabidopsis thaliana root gall showed actin and microtubules in a cell.

Concurrent immunolabelling for actin (A, B) and microtubules (C) in a cell in gall tissue. A Maximum projection of ten sections (sections 25-35) taken at 1 µm intervals. B, C A single optical sections at 30 µm depth. a, actin; mt, microtubules. Bar = 5 µm

2.3.2.5 GFP-hTalin to study actin in Arabidopsis thaliana root infected by Meloidogyne javanica

The GFP-hTalin Arabidopsis thaliana plants were ideal for studying root infection by M. javanica because GFP labelling of the actin cytoskeleton could be observed in the epidermal, cortical and vascular tissue, and changes caused by infection could be readily detected in comparison to uninfected roots.

Twenty-four hours after inoculation with Meloidogyne, actin filaments were mostly cortically distributed. Cells in close proximity to the nematode appeared to show more fluorescent labelling compared to cells further away from the nematode. Increased fluorescence was observed in and near nematode entry wounds (Figure 2.22).

Observation of Meloidogyne-inoculated roots after 48 h showed a nematode migrating within the root tissue. Epidermal cell shape in the roots was almost normal and actin filaments in these cells were more reticulate than in uninfected samples. There was increased actin labelling in cells close to the body of migrating nematode. In some cells actin filaments surrounded the nucleus of the cell and radiated to the outer cell wall during nematode migration, as it did in uninfected cells (Figure 2.23, arrows).

Figure 2.22 A GFP-hTalin Arabidopsis thaliana root 24h after inoculation showing entry by Meloidogyne javanica just behind the root tip.

GFP-hTalin fluorescence showed actin organisation (A-D) in a Meloidogyne javanica infected Arabidopsis thaliana root 24 h after inoculation. A Maximum projection of 26 sections. B-D Maximum projections of confocal optical images at 1-8, 9-17 and 18-26 respectively, taken at 1µm intervals from the surface of the root. E-G Concurrent transmitted light images at 8, 17 and 26 µm. Arrowheads point to a juvenile nematode which can be seen entering the root (outlined in white in E and F). Thin arrows show increased actin labelling in the gall tissue. Short arrow points to position of nematode in the root (A-D). Note the rounded shape of the gall cells. Bar = 50 µm.

Figure 2.23 A GFP-hTalin Arabidopsis thaliana root 48 h after inoculation showed Meloidogyne javanica juveniles migrating within the root.

GFP-h Talin fluorescence showed the actin organisation (A-D) in Arabidopsis thaliana 48 h after inoculation with Meloidogyne javanica. A Maximum projection of 45 sections taken at 1 µm intervals from the root surface. B-D Maximum projections of optical sections at 1-15, 16-35 and 36-45 µm respectively. E-G Transmitted light images at 15, 35 and 45µm. Short arrows point to the nematode within the root tissue which is outlined in white. Long arrows show increased actin labelling in cells adjacent to nematode body. Bar = 30 µm.

In more developed galls observed at 14 DAI, epidermal cells were rounded or spherical and unlike epidermal cells in uninfected roots. They were also smaller and more numerous and the actin filaments and bundles were mostly cortical (Figure 2.24). Soon after gall formation started, the roots became too thick for confocal observations of GFP-hTalin within galls.

Figure 2.24 GFP-hTalin Arabidopsis thaliana showed actin labelling in Meloidogyne javanica feeding sites in a 14-day-old root gall.

Confocal optical sections showed actin fluorescence (A-D) in a root gall induced by Meloidogyne javanica in Arabidopsis thaliana. A Maximum projection of 15 sections taken at 1 µm intervals from the root surface. B-D Maximum projections of sections taken at 1-5, 6-10 and 11-15 µm from the root surface respectively. E-G Single transmitted light images 5, 10 and 15 µm from the root surface. Long arrows point to the highly fluorescent gall tissue surrounding the nematode; the nematode is outlined in white and indicated by arrowheads (E-G); short arrow in E points to the approximate location of the head of the nematode. Bar = 50 µm.

2.3.3 Arabidopsis thaliana infected by Pratylenchus thornei

A. thaliana roots were inoculated with surface-sterilised P. thornei larvae. Immunolabelling 3 h after inoculation showed wounds on the root surface with a mass of amorphous material outside the wound that was labelled with actin antibodies (Figure 2.25). This mass was situated on the surface of the wound outside the affected cell. In neighbouring cells, actin was not disrupted.

Root tips examined 24 h after inoculation showed a well-defined round wound on the surface of the roots (Figure 2.26). This area retained the amorphous fluorescent labelling for actin on the surface in a disc shape, but distinct bundles and filaments were not present. Deeper sections showed a ring shaped structure surrounding the wound (Figure 2.26 D, I). It was not clear whether this structure originated from the plant or the nematode. Microtubules were disrupted or absent towards the surface (Figure 2.26 I). Normal cortical microtubules were seen in the cells adjacent to the wounded cell. Deeper sections of the wound area (Figure 2.26 J) showed cortical microtubules of the outer wall of a cell pressed close to the inner wall of the same cell forming a semicircular depression.

Pratylenchus were only observed close to the surface of the root, and never within roots during the first 24 h after inoculation. After 24 h, significant bacterial contamination developed which resulted in the death of both the host plant and the nematode. The source of the contamination was probably from bacteria that the nematodes carried in their gut before inoculation. Due to this contamination, studies could not be conducted using the surface-sterilised Pratylenchus thornei for longer than 24 h post-inoculation.

Figure 2.25 Immunolabelling of an Arabidopsis thaliana root tip 3 h after inoculation with Pratylenchus thornei shows the organisation of actin and microtubules.

Concurrent immunolabelling for actin (A-E) and microtubules (F-J) showed damage to root cells 3 h after infection by Pratylenchus thornei. Arrowheads show the location of the cytoskeletal damage. A,F Maximum projections of 12 sections taken at intervals of 1 µm from the root surface. B-E, G-J Single optical images at 2, 4, 7 and 12 µm from the root surface. Bar = 20 µm.

Figure 2.26 Immunolabelling for actin and microtubules in an Arabidopsis thaliana root 24 h after inoculation with Pratylenchus thornei showed that the cytoskeleton of the root epidermis was modified.

Concurrent immunolabelling for actin (A-E) and microtubules (F-J) of an Arabidopsis root tip 24 hours after inoculation with Pratylenchus thornei showed that actin and microtubules were affected differently by infection. A, F Maximum projections of 9 sections taken at 1 µm depth from the root surface. B-E, G-J Single optical images at 2, 3, 5 and 9 µm from the root surface. Arrowheads point to the epidermal damage. A-E Disc-shaped plug of material positively labelled for actin on the root surface near the wound from which Pratylenchus was probably feeding, before it was dislodged during processing. F-J Damage to microtubules in affected cells. Microtubules seem to be mostly absent from this area. Bar = 20 µm.

2.4 Discussion

The plant cytoskeleton plays a dramatic part in many plant functions, especially plant defense responses (reviewed by Wasteneys and Galway, 2003; Wasteneys and Yang, 2004; Takemoto and Hardham, 2003; 2004). Changes may occur in the host cytoskeleton either as a defense response or induced by the infection of the pathogen in the host cells. Infections by nematodes are no exception to this, with studies using GFP-transformed Arabidopsis plants showing cytoskeletal changes following nematode infection (Urwin et al., 1997; De Almeida Engler et al., 2004). The present study was undertaken to further investigate changes in cytoskeleton following nematode infections and compare them using three different investigative techniques.

2.4.1 Comparison of contrasting modes of infection using three different techniques

Collings and Wasteneys (2005) used double immunolabelling of whole fixed roots to study the plant cytoskeleton. A simple modification of this procedure was used for this study and was found to be greatly effective in observing the cytoskeleton in root tips and the differentiating region of nematode-infected and uninfected A. thaliana roots. This was a good technique to observe the cytoskeleton in roots but it was extremely difficult to obtain actin fixation using this method, especially in nematode- infected tissue, probably due to changes in the physiological characteristics of affected cells as compared to uninfected Arabidopsis cells for which the technique was originally developed. An alternative method for looking at fixed tissue involved sectioning samples embedded in BMM resin. Baskin et al. (1992) described an improved technique for studying microtubules by embedding and sectioning. This technique was slightly modified and used to compare infected and uninfected Arabidopsis and was found to be suitable to study both actin and microtubules. The advantages of this method were that it resulted in better preservation of actin structures in infected tissue. Sectioning also allowed observation within the gall, which was not possible with whole mounted roots, either fixed and immunolabelled, or living and expressing GFP. BMM sections proved that actin is not depolymerised in infected tissue, as had been reported by De Almeida Engler et al. (2004). On the contrary, well defined actin bundles were observed in infected tissue.

More recently Arabidopsis plants transformed with GFP fusion proteins have been used to study the in vivo distributions of the cytoskeleton (Kost et al., 1998; Ueda et al., 1999; Ueda and Matsuyama, 2000; Takemoto et al., 2003). These plants have been found suitable for studying Meloidogyne-infected roots of Arabidopsis (Urwin et al., 1997; De Almeida Engler et al., 2004). The GFP-hTalin-transformed Arabidopsis plants used in this study were suitable for studying changes in the actin cytoskeleton in Meloidogyne infected cells although the size of galls limited observations there. The advantage of this method was the short processing required before observation with the confocal microscope.

2.4.2 Nematode Entry

Actin microfilaments were found to react differently to microtubules in wounds caused by nematodes on the surface of root cells. Both Meloidogyne javanica and P. thornei were observed to destroy the Arabidopsis root epidermal cell wall while trying to enter the root, thus wounding the cell. Similar observations were made in Arabidopsis infected by Meloidogyne (Wyss et al., 1992) and in strawberry infected by Pratylenchus (Kurppa and Vrain, 1985). Plants inoculated with either Meloidogyne or Pratylenchus showed similar responses to initial wounding on the root surface by nematodes. Material labelled for actin seemed to accumulate in a rounded or disc-shaped plug on the root surface and seemed to line the wound surface on the inside of the cell. Similar observations have been made on the response to wounding of Xenopus oocytes by Bement et al. (1993; 1999). They also observed a distinct rounding of the wound prior to full healing and closure of a wound.

Wound plugs in plant hosts have been reported earlier in response to wounding (Aist, 1976). The actin cytoskeleton may react to mechanical wounding by nematodes or other means by oozing out with the cell contents and accumulating on the surface of the wound. It has been reported that initial changes in actin organisation in response to Nod factor application occurs in legume roots in 5 to10 min (Lhuissier et al., 2001). Microtubules in the same affected cells, on the other hand, seemed to hold their position and in doing so, they possibly provide structural stability to the forming cell wall. Hayano et al. (1988) made similar observations in marine algae

and suggested that microtubules may control cell shape. Although they held their shape in affected cells, physical disruption was observed where nematodes had tried to push their way into the root tissue. This initial response was first noted within 3 h of inoculation. The bright mass of green fluorescence on the root surface made it easy to spot entry points of Meloidogyne in confocal observations. Such observations have not been reported earlier.

Increased actin labelling was detected in nematode-infected root tissue using all three techniques used. This was especially prominent during initial entry and feeding site formation but not as noticeable during nematode migration. Similar observations were also made by De Almeida Engler et al. (2004). Intense GFP fluorescence has been observed earlier where Meloidogyne infections in plants were studied using Arabidopsis transformed with actin and microtubule binding proteins (Urwin et al., 1997; De Almeida Engler et al., 2004). De Almeida Engler et al. (2004) proposed that increased fluorescence may be due to the increase in tubulin and actin concentration in infected cells. However, with whole mounted and immunolabelled roots, the increase in labelling may also be due to nematode-induced changes in the cell walls resulting changes in permeability to primary or secondary antibodies and thus to labelling intensity of actin or tubulin. Temperature-dependant changes in cell wall architecture have earlier been reported in the rsw1 mutant of A. thaliana (Sugimoto et al., 2001). They suggested that in addition to reduction in cellulose synthesis, changes in alignment of microfibrils may weaken the cell wall. A restrictive temperature of 29º C resulted in noticeable increase in root diameter and reduction in the rate of elongation. At this temperature, root hairs which are not normally permeable to antibodies, become permeable. In the present study, diffuse fluorescence of microtubules and actin was always observed when fixation was not adequate in specimens.

2.4.3 Nematode Migration

Once nematodes entered the root, they caused minimum disturbance or disruption to tissue or cell files while migrating, first towards the root tip and then towards the differentiating tissue through the stele to find a suitable feeding site. Similar observations were made earlier (Sijmons et al., 1991; Wyss et al., 1992). The

nematode may be following physiological or chemical signals derived from the different cell types during different stages of its migration.

It was noted that microtubules in cells proximal to the nematode body showed a characteristic wavy form rather than the taut form in uninfected cells. This has not been reported before. The presence of wavy microtubules in host cells adjacent to the nematode may be due to the disruption or displacement of the cells caused by the nematode forcing its way through the root tip. In addition, there may have been some degree of tissue rearrangement in the host in relation to the nematode during fixation or other microscopy procedures (Dr David Collings, Personal Communication). Scientists have reported earlier, using immunolabelling techniques (Gravato Nobre et al., 1995) that within roots of Arabidopsis thaliana, Meloidogyne incognita juveniles loosen the middle lamella to enable intercortical migration. It is possible that some such chemical interaction caused microtubules to be loose and wavy in the cells observed. Wavy microtubules were induced in the Haemanthus endosperm by application of the microtubule-stabilising drug taxol (Bajer et al., 1982). Lead and copper were also found to induce wavy microtubules in garlic root tip cells (Liu et al., 2009). However, it is uncertain whether these results could explain the occurrence of wavy microtubules observed in my study. Wavy microtubules have also been observed in animal cells, in neurones (Tögel et al., 1998).

Conspicuous changes were noticed in optical properties of cell walls of Meloidogyne infected tissue as compared to uninfected tissue in inoculated plants. Wall ingrowths in Meloidogyne-infected giant cells have been reported earlier (Jones, 1976). Changes in optical properties observed in this study may be due to changes in chemical or physiological properties due to the effect of the nematode parasitism.

2.4.4 Nematode feeding sites

In Arabidopsis roots uninfected by Meloidogyne, cells in the elongating and differentiating region rarely divide (Dolan et al., 1993). Feeding sites induced by Meloidogyne were located well within the regions of the root which had completed differentiation. Notwithstanding the fact that cells in this region normally never divide, in Arabidopsis infected by Meloidogyne, several cells in the process of

division was observed. Host cells near the head region of the nematode enlarged in size forming “giant cells” soon after the nematode found a suitable feeding spot in the vascular region of the differentiating zone and became stationary. Within these “giant cells”, abnormal spindles and incomplete phragmoplasts were observed, but preprophase bands were absent. Similar observations were made by De Almeida Engler et al. (2004) in galls induced on A. thaliana by Meloidogyne incognita. They found that in giant cells, an actin and cortical microtubule cytoskeleton, although disturbed, was still visible. In addition, they observed a functional mitotic apparatus to be present that contained multiple large spindles and arrested phragmoplasts, but no pre-prophase bands. This lack of pre-prophase bands is also found in syncytial endosperm cells of Arabidopsis that contain mini phragmoplasts within them (Otegui and Staehelin 2000). The absence of a pre-prophase band in giant cells is an additional similarity to the normal endosperm in plants, which is detailed by Verma (2001). Giant cells have also been compared to transfer cells due to their similarity in wall structure (Jones and Northcote 1972; Jones 1976; Jones and Payne 1978; Jones and Gunning 1976; Jones and Dropkin 1976). This shows that such abnormal structures may occur in the plant naturally and the nematode only modifies this ability to suit its own purpose.

This study showed a well-defined, cortical and transversely-arranged layer of microtubules in cells in the gall through immunolabelling of whole fixed roots. Unfortunately, actin labelling was not successful in most cases in infected tissue. This might be due to inadequate fixation of actin. In immunolabelled BMM sections of infected roots, however, endoplasmic actin filaments and bundles were clearly observed in infection sites. This is in contrast to De Almeida Engler et al. (2004) who found that actin microfilaments failed to properly organize and appeared partially depolymerised throughout feeding site development.

Transverse arrangement of cortical microtubules is the main characteristic of actively growing, dividing and elongating cells (Yuan et al., 1994; Barlow and Baluška, 2000). In this study, microtubules within giant cells were found to be mainly transversely oriented. The nematode thus converted cells, which in the normal course of plant growth would have stopped growing and would be displaying oblique to longitudinal microtubules, to actively growing meristematic cells characterised by

transverse microtubules. The highly metabolically active nature of the giant cell was earlier described by Jones (1981a; b).

Uninfected Arabidopsis plants show tightly regulated growth polarity and growth anisotropy (Wasteneys, 2000). Studies using mutant alleles of RADIALLY SWOLLEN 4 and 7 found reduced growth anisotropy without altering the transverse orientation of cortical microtubules or cellulose microfibrils. In swollen regions of both mutants, cortical microtubules and cellulose microfibrils were neither depleted nor disoriented suggesting that factors additional to microtubules and microfibrils are necessary to limit radial expansion (Wiedemeier et al., 2002). However, recent studies by Bannigan et al. (2006) demonstrated that this original analysis was false, and that microtubules were disrupted in the rsw 6 mutant. Anisotropic expansion of the plant cell wall has been extensively reviewed by Baskin (2005).

Giant cells induced by Meloidogyne were irregularly shaped and seemed to have lost its growth polarity and anisotropy. The shape of giant cells observed resembled a long “spindle” with two tapering ends and an enlarged middle section. This may indicate that the factors necessary for expansion of the middle section of the giant cell may be ineffective in and therefore different from those needed for changing structure of the end walls of the cells. This is supported by the work of Collings and Wasteneys (2005) who found that increases in cell volume in the elongation zone resulted from anisotropic cell expansion with no obvious increase in cell diameter, implying that the cross walls have fundamentally distinct properties to the cell walls forming the main cell axis.

In several cases, nematodes were found to infect the root tip forming multiple galls. In these cases, cell files and even the quiescent centre were disrupted. Cell files of alternating trichoblasts and atrichoblasts were distinguishable even when there was extensive disruption of other tissue, and massive infection caused the root cap to break off in some cases. Atrichoblasts had more labelling of microtubules and actin filaments than trichoblasts. Similar observations were made by Collings and Wasteneys (2005). Root tip galls were observed earlier on Lotus japonicus (Lohar and Bird, 2003) which caused cessation of root growth from the meristem.

2.5 Conclusion

Changes in cytoskeleton were observed in root cells of Arabidopsis by Meloidogyne infection. Nematode entry caused unknown material labelled positively for actin to ooze out of the wounded cell causing a wound plug in infection by both Meloidogyne and Pratylenchus while the microtubule arrangement was mostly unaffected apart from the physical disruption. In feeding sites caused by Meloidogyne, extensive disruption of tissue, increased abnormal cell divisions and loss of polarity and growth anisotropy were noted. The three techniques used to compare effects on the cytoskeleton in infected roots were found to be very effective and can be used for further investigations.

CHAPTER 3

Effect of the nematodes Meloidogyne javanica and Pratylenchus thornei on growth and yield of tomato, Solanum lycopersicum

3.1 Introduction

The root-knot nematode, Meloidogyne javanica, and the root lesion nematode, P. thornei, are two of the most economically important nematode parasites in the world (see Chapter 1).

These two nematodes infect the tomato crop at all phenological stages of the plant. Not withstanding this fact, there is limited knowledge on the effect of infestation by these two nematodes in relation to different plant growth stages. There can be considerable variation in the population levels of each of these nematodes at which crop damage can occur. At very low populations, there may be very little effect on growth and yield of the host plant; at very high densities the plant can be severely damaged or may be even unable to recover from infection (reviewed extensively in Barker and Olthof, 1976). If it is possible to determine which stage(s) of the plant growth, when infected by these nematodes, results in the greatest loss in yield, this would assist in optimising timing for application of control measures. It would, in turn, facilitate a more sustainable approach to disease management, because targeting control measures at crucial stages may lead to more rational use of toxic chemicals and, in turn, less environmental pollution.

Nematode withdrawal of nutrients from plants at crucial stages of their growth may, in turn, affect their growth and yielding capacity. Up to 25% yield loss has been reported in pineapple in Queensland, even with a low population density of M. javanica of 50 per 100 g soil (Stirling and Kopittke, 2000).

In addition to direct damage by nematodes, secondary infection by other pathogenic organisms may occur through wounds on the root surface initiated by the nematode. For example, when tomato was treated concomitantly with Meloidogyne incognita

and Fusarium oxysporum f. sp. lycopersici it was found that shoot weight was reduced by 47% compared to 9% by Meloidogyne alone and 16% by Fusarium alone (Carter et. al., 1978).

Pratylenchus spp. is the most important species of plant parasitic nematodes involved in interactions with Verticillium wilt fungi. Either pathogen alone can cause the disease, but damage is greater when the nematode and fungus occur together (reviewed by Powell, 1971). Similar interactions have also been reported between Pratylenchus spp. and other fungal pathogens such as Pythium spp., Fusarium spp. and Rhizoctonia solani (Powell, 1971; Zuckerman et al., 1971).

3.2 Materials and Methods

Cultures of M. javanica and P. thornei were maintained in the laboratory at UWS according to the methodology previously described (Sections 2.2.1 and 2.2.2). These were used for inoculation, as outlined below.

As there was difficulty in accessing suitable facilities, the experiments with M. javanica and P. thornei had to be conducted separately, at different locations. Therefore, they were treated as two separate experiments; each with its own separate control.

Tomato plants, S. lycopersicum cv Grosse Lisse, were grown for one season from 20 November 2004 to 15 February 2005 in greenhouses at UWS, where they were used in the nematode experiments. Tomato seeds (total of 20 g) were germinated in two 30 cm diameter pots filled with potting mixture which comprised a blend of composted Pinus radiata pine bark (0 to 3 mm, fine and 7 to 10 mm, coarse), aged Pinus radiata pinewood (0 to 5 mm) and quartz sand (<2 mm). Sand provided the ballast and helped with wettability (minimum Australian Standards [AS 3743–2002] air-filled porosity of 13%) for germination of the seeds. At the four-leaf stage (including cotyledenous leaves), approximately two weeks after sowing, one tomato seedling each was transplanted into a 30 cm diameter pot filled with the previously described potting mixture. Slow release fertilizer (20 g, Osmocote®, Scotts Australia

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Pty Ltd, Baulkham Hills, NSW 2153) was added to each pot and mixed with the mixture just prior to transplanting.

Treatments consisted of inoculation with either M. javanica or P. thornei at planting and, thereafter, at weekly intervals up to nine weeks after planting, which were compared with a non-inoculated control. There were four replicates of each treatment. Nematodes were applied at the rate of 5000 per plant at a distance of 15 cm from the base of the stem of each plant in 1 mL sterile tap water, using a pipette. After an interval of 24 h after inoculation, watering to field capacity was applied once daily at 08:00. The experiment was conducted at ambient temperature, at a temperature of 25 ± 5° C (no specific temperature control was applied). Plants were checked regularly for pests and diseases.

One week after the application of the last nematode treatment, the plants were destructively sampled. This coincided with the first fruit harvest. The length of shoots, the number of green leaves, the number of flowers and flower buds and the number, fresh weight and diameter of fruits were recorded. Diameter of the fruit was taken from the largest mature fruit of each replicate at harvest. Then the shoot and root of each plant were separated by cutting through the stem at a distance of one cm from the point of emergence of the first root. The shoots, roots and fruit for each replicate were labelled and dried separately in a dehydrating oven (Thermoline Scientific, Smithfield NSW 2164) at 80° C for 48 h, and their dry weights were recorded using a Mettler Toledo PB 3002 DeltaRange balance (0.5 – 3100 ± 0.01 g, Precision Calibration Services, Wetherill Park, NSW 2164).

3.2.1 Statistical Analysis

Data were analysed using one factorial analysis of variance (ANOVA) general linear SPSS® for WindowsTM Version 12 (SPSS Inc. 2003). The assumption of normal distribution was checked using P-P plot and homogeneity of variance using Levene’s test of equality of error variances. Treatment means were tested against each other using Ryan’s Q test if the assumption of equality of variance was met or Dunnett’s T-test if the assumption was not met.

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3.3 Results

3.3.1 Observations on the growth stages of tomato, Solanum lycopersicum, in the greenhouse

The tomato plants reached the two-leaf and four- to five-leaf stages one and two weeks after sowing, respectively. Flowering commenced about six weeks from sowing (i.e. four to five weeks after transplanting). Flowering continued for approximately five weeks, and the fruits started to ripen one to two weeks after the start of flowering (viz. first ripe fruits were observed five to seven weeks after transplanting).

Galls and root lesions were clearly visible on all plants inoculated with M. javanica and P. thornei, respectively, at harvest.

3.3.2 Meloidogyne javanica

Shoot length Plants inoculated at planting and four and five weeks after planting had significantly shorter shoot length (F 10, 30 = 4.012, p = 0.001) than the uninoculated control plants (Table 3.1). Shoot length in plants inoculated one, two, three, six, seven, eight and nine weeks after planting did not differ significantly from that in the control. None of the nematode treatments differed from each other.

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Table 3.1: Summary of the statistical analysis for the mean shoot length, dry weight of shoot and the mean number of green leaves at harvest of tomato inoculated with Meloidogyne javanica (n=4) Treatment Mean shoot Significance Mean dry Significance Mean number Significance length (cm) (Dunnett’s T3 weight of (Ryan’s of green (Dunnett’s T3 (SE) Test) shoot (g) Q Test) leaves Test) (SE) (SE) No Nematodes 138.25 a 41.81 a 18.00 a (3.84) (2.75) (0.913) Inoculation at 102.00 b 20.85 b 10.75 a planting (3.39) (1.68) (1.75) Inoculation one 117.50 ab 29.40 b 10.75 a week after (4.91) (4.52) (2.562) planting Inoculation two 120.50 ab 21.73 b 7.25 a weeks after (5.60) (0.49) (2.02) planting Inoculation 105.25 ab 20.70 b 12.00 a three weeks (12.05) (4.73) (2.12) after planting Inoculation four 111.50 b 25.33 b 13.75 a weeks after (2.50) (2.25) (3.84) planting Inoculation five 97.75 b 25.48 b 12.75 a weeks after (5.51) (1.75) (1.89) planting Inoculation six 114.00 ab 25.48 b 14.50 a weeks after (4.24) (2.23) (2.47) planting Inoculation 111.75 ab 22.90 b 9.50 a seven weeks (3.57) (3.58) (3.34) after planting Inoculation 114.00 ab 25.03 b 11.00 a eight weeks (2.16) (2.52) (3.81) after planting Inoculation nine 115.75 ab 24.83 b 13.25 a weeks after (5.56) (2.68) (3.99) planting

Treatments with the same letter within the same column do not differ significantly from each other (p ≤ 0.05)

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Dry weight of shoot Plants inoculated at planting and thereafter at weekly intervals up to the ninth week after planting all had significantly lower dry shoot weight (F 10, 30 = 4.04, p = 0.001) than those in the uninoculated control (Table 3.1). However, the nematode treatments did not differ from each other.

Number of green leaves at harvest

There was no significant difference (F 10, 30 = 1.234, p = 0.310) in the number of green leaves at the time of harvest between plants inoculated at weekly intervals from the day of planting up to nine weeks after planting, and the uninoculated control, nor were there any differences between each other (Table 3.1).

Dry weight of roots Plants inoculated at planting and thereafter at weekly intervals up to nine weeks after planting had significantly lower dry weight of roots (F 10, 30 = 17.695, p = < 0.001) than the uninoculated control (Table 3.2). However, the nematode treatments were not significantly different from each other.

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Table 3.2: Summary of the statistical analysis for the mean dry weight of root and mean number of flowers and buds at harvest of tomato plants inoculated with Meloidogyne javanica (n=4)

Treatment Mean dry weight Significance Mean number Significance of root (g) (Ryan’s Q Test) of flowers and (Dunnett’s T3 (SE) buds at harvest Test) (SE) No Nematodes 51.75 a 4.25 a (2.45) (1.44)

Inoculation at 18.50 b 3.25 a planting (3.48) (1.97)

Inoculation one 21.00 b 8.50 a week after (1.87) (2.33) planting

Inoculation two 20.50 b 4.5 a weeks after (2.10) (1.76) planting

Inoculation three 18.75 b 3.1 a weeks after (3.54) (2.68) planting

Inoculation four 19.00 b 6.00 a weeks after (3.11) (2.97) planting

Inoculation five 20.50 b 4.50 a weeks after (1.76) (2.10) planting

Inoculation six 18.50 b 3.50 a weeks after (2.75) (0.20) planting

Inoculation seven 15.25 b 3.25 a weeks after (0.48) (1.89) planting

Inoculation eight 16.75 b 5.25 a weeks after (2.06) (1.70) planting

Inoculation nine 14.50 b 4.00 a weeks after (2.22) (1.23) planting

Treatments with the same letter in the same column do not differ significantly from each other. p ≤ 0.05).

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Total number of combined flowers and buds at harvest There were no significant differences in the combined number of flowers and flower buds on the plants at harvest (F 10, 33 =1.470, p = 0.195) between those inoculated at weekly intervals from the date of planting up to the ninth week after planting, and the uninoculated control, nor between any nematode treatment (Table 3.2)

Diameter of largest mature fruit Plants inoculated three weeks after planting had significantly lower mean mature fruit diameter (F 10, 30 = 2.222, p = 0.045) than those in the uninoculated control. However, plants inoculated at planting and one, two, four, five, six, seven, eight and nine weeks after planting did not differ significantly from the control (Table 3.3). There was no significant difference between any nematode treatments.

Total fresh weight of fruit at harvest Plants inoculated three weeks after planting had significantly lower fresh fruit weight

(F 10, 30 = 2.589, p = 0.021) than those in the uninoculated control. However they did not differ from any other nematode treatment. No other nematode treatment differed from the control. (Table 3.3).

Total dry weight of fruit Plants in all nematode inoculated treatments (from the date of planting to nine weeks after planting at weekly intervals) had significantly lower total dry weight of fruit (F

10, 30 = 4.60, p = 0.001) than control plants. There was no difference between any nematode treatment (Table 3.3).

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Table 3.3: Summary of the statistical analysis for the mean diameter of largest mature fruit, fresh weight of fruit at harvest, dry weight of fruit at harvest and moisture content of fruit at harvest of tomato inoculated with Meloidogyne javanica (n=4)

Treatment Mean diameter of Significance Mean fresh weight Significance Mean total dry Significance Mean moisture Significance largest mature (Ryan’s Q of fruit at harvest (Ryan’s Q weight of fruit (Ryan’s Q Test) content of fruit (F test) fruit (cm) Test) (g) Test) at harvest (g) at harvest (%) (SE) (SE) (SE) (SE) No Nematodes 7.10 a 272.25 a 24.67 a 90.93 a (0.14) (5.50) (0.98) (0.38) Inoculation at 4.93 ab 134.82 ab 13.75 b 89.69 a planting (0.68) (21.92) (2.17) (0.47) Inoculation one 5.95 ab 171.88 ab 13.59 b 91.57 a week after (0.35) (35.98) (1.81) (1.00) planting Inoculation two 5.50 ab 196.50 ab 15.60 b 91.79 a weeks after (0.44) (30.46) (1.83) (0.70) planting Inoculation three 3.67 b 102.78 b 11.90 b 89.16 a weeks after (1.26) (49.61) (1.76) (3.00) planting Inoculation four 4.65 ab 138.00 ab 12.73 b 90.55 a weeks after (0.16) (17.50) (0.94) (0.61) planting Inoculation five 5.68 ab 190.75 ab 14.83 b 91.92 a weeks after (0.21) (30.48) (1.60) (0.66) planting Inoculation six 5.68 ab 166.94 ab 14.78 b 90.92 a weeks after (0.82) (32.74) (2.24) (0.53) planting Inoculation seven 4.78 ab 169.25 ab 13.43 b 91.94 a weeks after (0.46) (15.64) (0.59) (0.50) planting Inoculation eight 5.73 ab 167.35 ab 14.68 b 90.87 a weeks after (1.07) (29.89) (1.91) (0.71) planting Inoculation nine 6.60 ab 230.60 ab 16.50 b 92.63 a weeks after (0.45) (29.58) (0.91) (0.58) planting

Treatments with the same letter in the same column do not differ significantly from each other (p ≤ 0.05). 107

Mean moisture content of fruit

The mean moisture content of fruits of plants inoculated with M. javanica varied between 89.2% for plants inoculated three weeks after planting to 92.6% for plants inoculated nine weeks after planting. This was similar to the moisture content of the control plants (90.9%). There was no significant difference between any treatment (F

10, 32 = 1.261, p = 0.293) (Table 3.3).

3.3.3 Pratylenchus thornei

Shoot length Plants inoculated at planting and one, two, three, five six, seven, eight and nine weeks thereafter, had significantly shorter shoot length (F 10, 30 = 4.415, p = 0.001) than plants in the uninoculated control. However, shoot length did not differ significantly between the control and plants inoculated at four weeks after planting (Table 3.4). There were no differences in shoot length between any of the nematode treatments.

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Table 3.4: Summary of the statistical analysis for the mean shoot length, dry weight of shoot and mean number of green leaves of tomato inoculated with Pratylenchus thornei (n=4)

Treatment Mean shoot Significance Mean dry weight of Significance Mean number of green Significance length (cm) (Ryan’s Q shoot (g) (Ryan’s Q test) leaves (Ryan’s Q Test) (SE) Test) (SE) (SE) No Nematodes 141.25 a 24.43 a 14.00 a (11.34) (1.07) (0.91) Inoculation at planting 98.25 b 13.40 b 9.50 ab (7.40) (0.85) (0.65) Inoculation one week after 84.75 b 16.16 b 9.50 ab planting (5.92) (0.74) (1.76) Inoculation two weeks after 100.50 b 16.88 b 9.25 ab planting (6.76) (1.06) (1.32) Inoculation three weeks after 101.75 b 15.66 b 7.25 b planting (8.29) (0.73) (1.38) Inoculation four weeks after 115.00 ab 15.38 b 9.50 ab planting (7.71) (1.37) (1.56) Inoculation five weeks after 92.00 b 17.86 b 11.00 ab planting (2.97) (1.44) (1.56) Inoculation six weeks after 102.25 b 17.93 b 9.75 ab planting (4.91) (0.63) (0.75) Inoculation seven weeks after 95.75 b 16.06 b 8.75 ab planting (7.59) (1.36) (1.32) Inoculation eight weeks after 101.50 b 18.09 b 9.00 ab planting (2.84) (0.83) (1.23) Inoculation nine weeks after 97.25 b 16.08 b 5.75 b planting (5.36) (0.68) (1.45)

Treatments with the same letter in the same column do not differ significantly from each other (p ≤ 0.05).

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Dry weight of shoot All nematode-inoculated tomato plants had significantly lower dry shoot weights (F

10, 30 = 7.434, p = < 0.001) than uninoculated control plants, but did not differ significantly from each other (Table 3.4).

Number of green leaves at harvest Tomato plants inoculated three and nine weeks after planting had significantly fewer green leaves at harvest (F 10, 30 = 2.512, p= 0.025) than plants from the uninoculated control. However, plants inoculated at planting and one, two, three, four, five, six, seven and eight weeks after planting did not differ from those in the control (Table 3.4). There were no differences between any of the nematode treatments.

Dry weight of roots Plants inoculated at weekly intervals from the date of planting to nine weeks after planting did not differ significantly from the uninoculated control (F 10, 30 = 1.305, p = 0.273) with respect to dry root weight (Table 3.5).

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Table 3.5: Summary of the statistical analysis for the mean dry weight of root and number of flowers and buds at harvest of tomato inoculated with Pratylenchus thornei (n=4)

Treatment Mean dry weight Significance Mean number of Significance of root (g) (Ryan’s Q Test) combined flowers (Dunnett’s T3 (SE) and buds at harvest Test) (SE) No Nematodes 50.75 a 8.5 a (10.52) (0.5) Inoculation at 35.75 a 6.75 ab planting (1.97) (0.85) Inoculation one 47.75 a 2.25 b week after planting (8.92) (0.63) Inoculation two 41.50 a 2.50 b weeks after (4.41) (0.50) planting Inoculation three 36.25 a 2.75 b weeks after (7.70) (0.25) planting Inoculation four 35.00 a 3.50 b weeks after (2.48) (1.56) planting Inoculation five 36.00 a 4.50 ab weeks after (2.83) (1.71) planting Inoculation six 39.50 a 4.0 ab weeks after (3.95) (1.35) planting Inoculation seven 33.00 a 2.50 b weeks after (2.42) planting (1.56) Inoculation eight 44.25 a 1.75 b weeks after (2.25) (0.25) planting Inoculation nine 35.75 a 2.25 b weeks after (1.97) (1.32) planting

Treatments with the same letter in the same column do not differ significantly from each other (p ≤ 0.05).

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Total number of combined flowers and buds at harvest Plants inoculated one, two, three, four, seven, eight and nine weeks after planting had significantly fewer combined flower and flower bud numbers (F 10, 33 = 3.78, p = 0.002) compared to the uninoculated control (Table 3.5). However, plants inoculated at planting, and five and six weeks after planting did not differ significantly from the control. There were no differences between any of the nematode treatments.

Diameter of largest mature fruit Plants inoculated from the date of planting to nine weeks after planting at weekly intervals did not differ significantly from each other nor from the uninoculated control (F 10, 30 = 1.536, p = 0.175) with respect to the diameter of the largest mature fruit at harvest (Table 3.6).

Total fresh weight of fruit at harvest There was no significant difference in total fresh fruit weight between any nematode treatment, nor between treatments and the uninoculated control (F 10, 33 = 0.511 p = 0.870) (Table 3.6). .

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Table 3.6: Summary of the statistical analysis for the mean diameter of largest mature fruit, fresh weight of fruit at harvest, dry weight of fruit at harvest and moisture content of fruit at harvest of tomato inoculated with Pratylenchus thornei (n=4)

Treatment Mean diameter Mean fresh Mean dry Mean moisture of largest weight of fruit weight of fruit content of fruit mature fruit at harvest (g) at harvest (g) at harvest (%) (cm) (SE) (SE) (SE) (SE) No Nematodes 6.31 59.44 4.97 91.46 (0.12) (6.67) (0.34) (0.58) Inoculation at 4.35 50.51 5.34 87.67 planting (0.29) (19.66) (1.69) (2.42) Inoculation one 4.64 51.88 7.71 85.09 week after (0.52) (5.84) (0.81) (0.23) planting Inoculation two 4.69 65.81 8.29 84.67 weeks after (0.91) (29.81) (2.24) (2.18) planting Inoculation 5.38 74.45 9.31 87.49 three weeks (0.38) (8.17) (1.76) (1.87) after planting Inoculation four 4.15 50.25 7.20 84.36 weeks after (0.25) (12.98) (2.12) (3.02) planting Inoculation five 5.05 60.38 9.72 83.92 weeks after (0.41) (7.09) (1.28) (0.68) planting Inoculation six 4.90 65.76 8.48 87.21 weeks after (0.35) (12.61) (1.91) (2.24) planting Inoculation 5.55 72.12 8.97 87.37 seven weeks (0.85) (20.72) (3.44) (2.26) after planting Inoculation 4.95 54.69 7.40 85.97 eight weeks (0.48) (10.68) (1.39) (1.57) after planting Inoculation nine 5.60 86.56 11.43 86.63 weeks after (0.37) (22.21) (2.73) (0.63) planting

There was no significant difference between any treatments (p ≤ 0.05).

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Total dry weight of fruit

There was no significant difference (F 10, 33 = 0.913, p = 0.533) in the total dry weight of the fruit at harvest between plants inoculated at weekly intervals from the day of planting up to nine weeks after planting, and the uninoculated control, nor between each other (Table 3.6).

Mean moisture content of fruit

The mean moisture content of fruits in plants inoculated with P. thornei varied between 83.9% for plants inoculated five weeks after planting to 87.7% for plants inoculated at planting. These were all less than the moisture content for control fruit

(91.5%). However, there was no significant difference between these data (F 10, 33 = 1.456, p= 0.200)

3.3.4 Comparison of growth and yield of tomato, Solanum lycopersicum, inoculated with Meloidogyne javanica and Pratylenchus thornei

The effects of the two nematode species on tomato growth and developmental parameters were compared for each nematode species using percentage change from control and combined nematode inoculation times, using the formula:

c − t X 100 c where c is the mean control value and t the mean treatment value.

The results are presented in Table 3.7.

Dry weight of shoots Both species of nematodes reduced the dry weight of shoots in tomato plants. However plants inoculated with M. javanica had an overall higher percentage reduction in dry weight of shoot (41.7%) than plants inoculated with P. thornei (33.1%) (Table 3.7).

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Table 3.7: Percentage reduction of the dry weights of shoot, root and fruit, number of combined flowers and flower buds and fresh weight of fruit, from control

Nematode Dry weight of Dry weight of Mean number Fresh weight Dry weight of species shoots roots of combined of fruit fruit (SE) (SE) flowers and (SE) (SE) flower buds (SE) M. javanica 41.73 64.13 -9.80 37.13 42.28 (2.17) (1.45) (14.70) (3.46) (2.09)

P. thornei 33.08 24.19 61.47 50.76 24.35 (1.45) (2.85) (4.56) (3.13) (4.27)

Dry weight of roots The percentage reduction in dry weight of roots of plants inoculated with M. javanica was greater (64.1%) than that for plants inoculated with P. thornei (24.2%) (Table 3.7).

Total number of combined flowers and buds at harvest Plants inoculated with M. javanica had a slightly higher number of flowers and buds at harvest (9.8%) than the uninoculated control, whereas plants inoculated with P. thornei had 61.5% fewer flowers and buds than their respective control plants (Table 3.7).

Total fresh weight of fruit Plants inoculated with M. javanica had 37.1% lower fresh weight of fruit from plants than control plants, whereas plants inoculated with P. thornei had 50.8% lower fresh weight of fruit than their control (Table 3.7).

Total dry weight of fruit

Plants inoculated with M. javanica had 42.3% lower dry weight of fruit than control plants. Plants inoculated with P. thornei had only 24.4% lower dry weight of fruit, compared to their control (Table 3.7).

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3.4 Discussion

Crop loss studies have been conducted in several field crops over the past few decades. Most of the studies arrived at their results by applying nematicides or other control methods to crops and comparing the yield of plants in which no control was applied. My experiment was aimed at comparing the effect of nematodes on tomato growth and yield when exposed at its different growth stages.

The tomato plants used in my experiment, S. lycopersicum var. Grosse Lisse, reached the two-leaf and four to five leaf stages one and two weeks after sowing, respectively. Flowering commenced about six weeks from sowing (four to five weeks after transplanting). Flowering continued for approximately five weeks, and the fruits started to ripen one to two weeks after the start of flowering. The first ripe fruits were observed five to seven weeks after transplanting, and the plants, including fruits, were harvested at ten weeks after transplanting. Tomato is known to be a photo- and temperature- sensitive plant (Aung and Austin 1971), with its growth best between 20 to 27° C (Benton Jones, 2007). Hussey (1965) reported that the optimum day temperature for dry matter deposition was 25 °C. My experiment was conducted in ambient temperature during the summer of November 2004 to February, 2004 when the temperature range was 25 ± 5° C. The growth and development of plants were not adversely affected at this temperature. This temperature was also suitable for both nematode species used in this experiment (see Chapter 1).

Tomato passes through different phenological stages over the course of its lifetime. There are several different classifications of these phenological stages, even up to 21 classes in some cases (Zalom and Wilson, 1999). According to Benton Jones (2007), the approximate time from planting to market maturity for an early variety is from 50-65 d while for a late variety it is from 85-95 d. The time period from seeding to first fruit harvest will vary with the maturity class of the cultivar from as little as 45- 100 d, and can approximately be divided thus:

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Table 3.8: Developmental stages and their durations in tomato, Solanum lycopersicum

Stage Number Developmental Stage Length Stage (d) 1 Establishment 25-35 (germination to initial leaves)

2 Vegetative 20-25

3 Flowering 20-30

4 Fruit formation 20-30

5 Fruit ripening 15-20

Total 100-140

For the purposes of this experiment, growth of tomato consisted of mainly three stages, vegetative stage, flowering stage and fruiting stage.

There is no standardised level of nematode inoculum level and stage for field or pot investigations, and previous studies have used widely varying initial inoculum densities in assessing crop losses. This makes comparisons between different studies difficult. Di Vito et al. (1986) found that inoculum type affected experimental results on eggplants. They demonstrated that intact M. incognita egg masses gave rise to more juveniles than from eggs which had been released from the gelatinous matrix by hypochlorite. Also, the infectivity of juveniles from intact egg masses was greater. Using chopped, infested roots as an inoculum was apparently even more effective, because plant yields were reduced even further.

Inoculation with one J2 M. incognita per gram of soil caused a significant reduction in growth of eggplants (Dhawan and Sethi 1976). In another study, an initial population density of 4400 M. incognita per kg in the top 30 cm. of soil reduced tomato yield significantly (Johnson and McKeen, 1973). Similar population densities at lower soil depths (120 to 135 cm from the surface) reduced tomato yield by 11%

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in the first crop and 59% in the second. A much higher population level of up to 20,000 Meloidogyne race 1 per plant was required to reduce the top weight of sugarcane in Brazil (Regis and Moura, 1989), and an initial inoculum level of 6000 eggs of M. incognita or M. javanica in bean, P. vulgaris, showed no effect on plant growth or nitrogen levels (Moura and Moura, 1994).

Haseeb et al. (1998) observed a negative correlation between various inoculum densities of M. incognita and fresh/dry weight, oil yield, photosynthetic rate, total chlorophyll, sugars, phenols and root-knot development in Ocimum kilimandscharicum. For assessing nematode resistance in tomato, 200 eggs or infective juveniles of Meloidogyne incognita or M. javanica per seedling was found to be the optimum initial inoculum level (Araujo et. al., 1982).

In the present study using 5000 nematodes per plant, it was found that, irrespective of the growth stage at which nematodes were applied, some growth parameters were adversely affected. Inoculation with M. javanica reduced the shoot length, diameter of fruit, dry weights of shoots and roots and the fresh and dry weights of fruits, while it did not affect the number of green leaves on the plant or the number of combined flowers and buds. Inoculation with P. thornei adversely affected the shoot length, number of green leaves, number of combined flowers and buds and the dry weights of shoots and roots. P. thornei did not affect fresh weight of fruit, diameter of the largest mature fruit and dry weight of fruit at harvest.

Vaast et al (1998) found that the uptake of nitrate and ammonia was greatly reduced by infections of Meloidogyne and Pratylenchus on coffee. When nematodes withdraw nutrients from the host for its growth and reproduction at a time of active growth, the host may adjust its growth to maximise its survival with the reduced food and water available to it. This is likely to result in reduction in growth parameters such as those I assessed in my study.

One of the aims of the investigation was to determine which stage of the plant growth, when infected by these nematodes, resulted in the greatest yield loss. My results suggest that even a short period of exposure to nematodes can adversely affect plant growth and development. For example, even two weeks’ exposure to P. thornei

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caused significant reduction in shoot length and dry weight of shoot in tomato. A reason for this may be that the plants were still relatively young; they had just commenced flowering and fruiting, so the effects on these parameters may still be regarded as due to infestation of young plants. The results may have been different with more mature plants. There is evidence that if mature plants are infested with nematodes, yield reduction is less significant (Khan, 2000) A second reason may be that plants exposed to nematodes earlier (viz. inoculation at planting or one or two weeks after planting) may have been able better compensate by, for example, initiation of new roots. Such compensation has been previously reported (Khan, 2000).

Reduction in the growth parameters recorded in my investigation was in contrast to observation by Antonio and Dall’Agnol (1982) who reported that dry matter production in soybean was not significantly affected by inoculum levels. Wallace (1973) reported that low infection levels stimulate plant growth whereas high nematode densities suppress plant growth and yield. In this crop loss study 5000 M. javanica and P. thornei per pot was used for inoculation, and this level of inoculum caused reduction in some growth parameters. However, further studies should be conducted to determine whether this inoculum level is the most appropriate to assess nematode effects on all plant growth characteristics. Further, the growing medium for this experiment was a uniform potting mixture (details in Section 3.2) and plants were grown in pots, which may not be directly applicable to field situations. In the field there are variable soil types with different textures, physical and chemical characteristics. However, I still expect that results obtained from this study would be helpful in understanding field situations of plant attack by nematodes. I have addressed the issue of soil type and movement of nematodes in the following chapter.

3.4.1 Effect of Meloidogyne javanica on growth and yield of Solanum lycopersicum

When tomato plants were inoculated with M. javanica four or five weeks after planting, the shoot length was reduced significantly. Inoculation with Meloidogyne three weeks after planting caused significant reduction in the diameter of the largest

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mature fruit at harvest. There were significant reductions in dry weights of shoot and root in inoculated tomato irrespective of the timing of inoculation. These results suggest that the third, fourth and fifth week after planting are critical stages in the growth of tomato. This may be due to withdrawal of nutrients essential to growth by the Meloidogyne for its own growth and reproduction, resulting in less food and water being available to the tomato plants. As first flowering was noticed at five weeks after planting, being deprived of essential nutrients and photosynthates in the weeks just prior may have affected growth and yield. However, infection by this nematode is unique in the way that it alters the physiology of the host and remains unchallenged within the host tissue, feeding on it and completing its life cycle without triggering major defense responses. Thus the very mode of infection by this nematode may have resulted in there being no significant effect on some other growth parameters such as the number of green leaves, the number of combined flowers and buds and the fresh and dry weights of fruit.

The percentage of moisture content in fruit in Meloidogyne-inoculated tomato ranged from 89.8% to 92.5% while that in the uninoculated control was 91.1%. This appears to show that water uptake of young tomato plants plant was not significantly reduced when they were infected by M. javanica. This is consistent with observations described in Chapter 5, where wound xylem was seen to be differentiating around nematode-disrupted xylem within a short period of time. This further suggests that the physiology of the interaction between the host (tomato) and the nematode (M. javanica) has a major role in the tomato plant growth and development, in addition to any direct physical damage caused.

3.4.2 Effect of Pratylenchus thornei on growth and yield of Solanum lycopersicum

Inoculation with P. thornei affected several growth parameters of tomato plants, including the shoot length, number of green leaves and the number of flowers and buds at harvest.

The dry weight of shoots and roots was also significantly reduced in Pratylenchus- inoculated plants. This result is supported by Castillo et al. (1998) who reported

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significant reduction in dry shoot and fresh root weight in Cicer arietinum caused by P. thornei. However, the fresh and dry weights of fruit and the diameter of the largest mature fruit were not affected by inoculation of plants with P. thornei.

Interestingly, the fruits of plants inoculated with P. thornei had a consistently lower moisture content (range 83.9 - 89.4%) than that recorded in the uninoculated control (91.6%). This latter figure is similar to the control and treatment plants for the M. javanica experiment. This difference in moisture content may be caused by the physical destruction to the root system associated with feeding and migration of the nematode within the root, which is likely to directly impact on water translocation from the soil to the other parts of the plant, including the fruits.

3.4.3 Comparison between effects of Meloidogyne and Pratylenchus on growth and yield of Solanum lycopersicum

The comparison between the effects of the two nematode species on key growth and development parameters provided some interesting results. For instance, there was a small increase in flowers and buds in Meloidogyne treatments compared to untreated control plants, but 60% reduction in Pratylenchus treatments compared to the control. It may be that infection by Meloidogyne could have induced an earlier flowering flush, although this has not been previously reported. The impact of Pratylenchus in reducing flowering is likely to be a direct response by the plant to root damage, which is likely to reduce plant vigour, or delay flowering. For all dry weight parameters, namely dry shoot weight, dry root weight and dry total fruit weight, Meloidogyne appeared to induce a greater reduction than Pratylenchus. However, this was reversed in the case of total fresh fruit weight. This phenomenon may be explained by the differences in fruit moisture content in Meloidogyne- infested plants (ranging from 89.8% to 92.5%), as compared to Pratylenchus-infested plants (ranging from 83.9% to 89.4%). In turn, this difference in moisture content is likely to be a consequence of the way in which these two nematode species attack and damage plant roots. This matter will be discussed in more detail in the final chapter. The general reduction in growth and development recorded here may be due to reduction in the efficiency of water and nutrient uptake by the nematode-damaged roots, or by changes in root physiology induced by the nematodes after infection.

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Significant reductions in uptake of ammonium and nitrate have been reported in coffee infected by Meloidogyne and Pratylenchus (Vaast et al., 1998).

3.5 Conclusions

M. javanica and P. thornei significantly affected selected growth parameters of tomato. M. javanica reduced shoot length and diameter of largest mature fruit growth mainly in the range of three, four and five weeks after planting. On the other hand, P. thornei reduced shoot length, dry weight of shoots, the number of green leaves, dry weight of roots, and the combined flowers and flower buds at least at some time between planting and nine weeks after planting. This is likely to be a critical period when tomato seedlings are rapidly growing, and are also starting their reproductive phase. Flowering in tomato was observed from the sixth week after planting onwards. Redirecting their resources towards reproduction, combined with the draining of nutrients by the nematode for its own growth and reproduction, probably resulted in adverse effects on the growth and yield of the tomato plants. It was interesting to note that even one week exposure to both nematode species resulted in a significantly lower values in a number of the parameters measured.

The significantly lower dry weights of shoot root and fruit in plants inoculated with Meloidogyne compared to plants inoculated by Pratylenchus indicates that the growth and physiology of tomato plants are influenced in different ways by the species of nematode involved. The contrasting modes of parasitism by these nematode species may in some way affect plant growth parameters. M. incognita changes the physiology of plant tissue so as to remain undetected as an enemy and manipulates the host, inducing formation of feeding sites and galls. P. thornei is mainly an indiscriminate, destructive feeder. It is possible that the physiological changes induced by Meloidogyne, which included rapid repair of the disrupted xylem tissue observed in this study (see Chapters 2 and 5), probably resulted in normal or near-normal moisture levels of moisture in fruit (i.e. as compared to the control plants) but production of less dry matter. The destruction of root tissue by the migration and feeding of Pratylenchus (see Chapter 5) may have affected the uptake of water and nutrients by plants parasitised by the nematode. This issue will be discussed in more detail later in this thesis.

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Results of this study suggest that if nematode control can be targeted at the vegetative phase, for example from transplanting to nine weeks after transplanting, it is likely to benefit growth and development in tomato. My studies did not involve nematode inoculations any later than this time, so it may be that growth and development may be affected even in the mid-reproductive phase. Finding the right time for applying control strategies will also result in savings in the cost of pest control, and increase crop yield, thus benefitting the primary producer. Thus, the results of this study will help to formulate integrated pest management methods suitable to obtain the most efficient use of resources such as biological control methods, alone or in combination, thus reducing the quantity of harmful pesticides used in crop cultivation.

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CHAPTER 4

Movement of Meloidogyne javanica and Pratylenchus thornei through sand, sandy clay loam and clay soils to the roots of tomato, Solanum lycopersicum

4.1 Introduction

Soil is the major habitat of nematodes (Koltai et al., 2001). Most plant parasitic nematodes spend at least part of their life cycle in soil. Sensory organs of plant parasitic nematodes enable them to find a suitable host in soil by responding to host and environmental stimuli like gradients of temperature, root exudates (Prot, 1980), and soil properties such as texture, composition, pore size and water holding capacity (Wallace, 1958; 1960). Very few studies have been published on influence of soil on nematode movement and most of this research was conducted in the latter half of the 19th century. Results of the above-mentioned studies suggest that physical and chemical factors in the soil and morphology and physiology of soil-dwelling nematodes can have important effects on their movement and consequent pathogenicity.

4.1.1 Soil types and soil texture

Soil is a naturally occurring mixture of mineral and organic matter on the surface of the earth. There are big variations in soils from location to location all over the world. However, soils may also vary even within a farmer’s field. Several factors influence this variation such as the parent material from which it was formed and its degree of weathering, topography, organisms (both large and small) in and around it, and climate including the amount of precipitation. In nature, soil is mostly composed of minerals, organic matter, air and water.

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Soil texture is an important physical property which is based on the relative proportions of its different sized mineral particles, namely sand, silt and clay. The particle sizes for these categories varies between countries; the most commonly used is that of USDA, where sand particles are the largest, between 2.00 - 0.05 mm, silt is 0.05-0.002 mm, and clay is < 0.002 mm. The relative proportions of these three particles have a major impact on a number of physical characteristics of the soil, including porosity, aeration, drainage and water-holding capacity. There are a number (11 in Australia. 12 in USA) of soil texture classes, which are determined by plotting on a 3-axis graph, known as a soil texture triangle (Minasny and McBratney, 2001).

Sandy soils in nature tend to be low in fertility, low in their ability to retain nutrients and moisture, and rapidly permeable. This is because of the large mean pore sizes, and the lower surface area to volume ratio of the particles. The relative proportion of silt and/or clay particles increases they naturally tend to be more fertile, have higher cation exchange capacities and are better able to retain moisture and nutrients. Soils with relatively even proportions of sand, silt, and clay are known as loams or “loamy”. Sand and silt are relatively inert, but clay is the active part- attracting and holding water and nutrients. However, because of the relatively higher number of small pores, they also tend to reduce movement of air and water (Brown 2003).

4.1.2 Organic matter in soils

Organic matter in soil comes from live as well as decomposed plant, animal and microbial components. It binds mineral particles into aggregates, strongly influences the manageability and productivity of soils, and also improves its water holding capacity. Partially decomposed organic matter is called humus, and is a good source of nutrients when combined with colloidal clay particles. Soils rich in organic matter have been shown in earlier studies to reduce crop losses caused by plant parasitic nematodes (Sikora et al., 2005; Widmer et al., 2002; Ciancio and Mukerji 2007).

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4.1.3 Pore space and aeration in soils

An ideal agricultural soil consists of 50% solid space and 50% pore space. Solid space consists of mineral and organic matter, and pore space contains water and air. Ideally pore spaces are half filled with air and half with water. Pore space plays a very important role in soil aggregation, nutrition and water holding capacity. Pores are the result of the irregular shape of soil particles, and vary with soil texture. Sand soils have smaller amounts of total pore space, but larger sized pores. Clay soils have more total pore space, but smaller sized pores. When pore space is very small, soil retains water, and drainage is impeded, resulting in poor aeration. Sufficient soil aeration is necessary for optimum plant growth and yield. Plant roots require oxygen to respire, releasing carbon dioxide into the soil. Soil microorganisms also release carbon dioxide into the soil during respiration. If there is inadequate aeration, this carbon dioxide accumulates in soil, depleting the available oxygen, which is detrimental to plant health. Usually, coarse, well drained soils have high oxygen content and low carbon dioxide content and fine textured soils have a high carbon dioxide and low oxygen content. If the proportion of clay particles in sandy clay loam soils is high, they can clog the pore spaces thus affecting drainage and reducing aeration. Clay soils may become waterlogged because of impeded drainage. Soil water contains dissolved nutrients which are available to soil inhabitants, and also decides the pH of the soil. Nematodes require a film of water to facilitate their movement through soil. But too much moisture is detrimental to the nematode population. High populations of nematodes are usually found in moist, well aerated soils.

Three types of soil (sand, sandy clay loam and clay) were used in this study to investigate the effects of soil characteristics on the movement of Meloidogyne javanica and Pratylenchus thornei to the roots of tomato.

4.1.4 Nematode morphology

An important factor which may influence the nematode movement towards the host root is the nematode morphology, or the size and shape of its body.

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A typical M. javanica second-stage larva, which is the only infective stage in its life cycle, is about 402-560 µm in body length (Eisenback and Triantaphillou, 1991) and about 10-15 µm in width.

P. thornei varies in length from 0.45 to 0.77 mm (Loof, 1991) and the body diameter is approximately 35 µm wide. Pratylenchus is at least three times as wide as M. javanica and body length at least 50% longer; it is, thus, a larger and stouter nematode in its infective stage than Meloidogyne.

4.1.5 Relationship between soil and movement of nematodes

There have been a few previous studies to investigate the optimum soil particle size for nematode movement.

Optimum particle sizes for movement of potato cyst nematode, Heterodera rostochiensis, in a sandy loam, heavy clay and peat soil were 150-250 and 250-400 µm (Wallace, 1958). Mobility was very similar in clay and sandy loam. In peat, mobility increased with suction pressure from 0 to 100 cm, and then decreased. Larvae moved to the wet end of a moisture gradient in sand. The rate of spread of larvae in sand 150-250 µm diameter varied between 2 and 3 cm per day, depending on suction. As pore size increased, any upward movement was opposed by effects of gravity. Nematodes did not respond to a moisture gradient or fall under gravity in sand where the width of the pore approximated the diameter of the larva. Mobility was greatest when (a) there were few pores smaller in diameter than nematode width, (b) the pore diameter was narrow enough to restrict lateral movement, (c) the channels between particles was such that the body form of the nematode had waves of long wave-length and short amplitude.

Different genera of nematodes prefer different types of soil for infection. Some studies have found that Meloidogyne prefers sandy soils for maximum galling (Verma and Jain, 1998; Desaeger and Rao, 2000). In a study conducted in French West Indies (Cadet and Thioulouse, 1998) Pratylenchus coffeae preferred any type of soil rich in organic matter. But it was not clear whether the authors meant preference of the nematode for survival in soil or for damaging the host in these

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types of soil. Recent studies showed that apparently small differences in soil composition between fields located in the same climatic area and managed similarly can translate into contrasted nematode damage levels and sugarcane yields (Rime et al., 2003). Prot and van Gundy (1981) suggested that clay particles aid in the migration of root-knot juveniles over long distances to plant roots by absorbing and holding root exudates or bacterial by-products which form a concentration gradient enabling nematodes to locate roots. Earlier studies (Prot, 1980) showed that juveniles of Meloidogyne were able to travel relatively great distances before infecting a susceptible tomato and that larvae preferentially moved to the region having more moisture and lower salt concentration.

Other studies have shown that nematodes prefer sandy loam soils to the host roots under certain conditions. Studies on Xiphinema index (Sultan and Ferris, 1991) showed that in the presence of a host, population increase of the nematode was highest in sandy loam and in fine sands of 250 µm particle size. Population increase was low in coarse sand of particles 534 µm and larger. They also found that root damage to host plants was directly related to the increase in nematode populations.

The presence of host roots can counteract the response to a moisture gradient; the degree of orientation to the roots increased with the time the roots were in the sand. Direct observation of the larvae, newly emerged from the cysts, in the presence of host plant roots, suggested that larvae orientated themselves at a distance from the root and did not reach the root by random movement (Wallace, 1960).

Other studies found that populations of Meloidogyne from West Africa have the capacity to migrate relatively long distances. Even though populations vary, it could be assumed that the same ability exists within Meloidogyne population in other geographical areas as well (Prot, 1980). Some nematodes are able to migrate over long distances, horizontally and vertically, and some are not. It is less probable that temperature, redox potential or CO2 created by the roots have any effect on migration over long distances as compared to short distances from 1 to 2 cm. In this case only the secretion of roots or their bacterial by products are more likely if they are carried by water percolation (Prot, 1980; Prot and van Gundy, 1981).

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Thus, many of the movements of nematodes are explicable by considering the relationship between pore size, soil texture type, water distribution and, possibly, organic matter content. Considering the fact that soil properties play such an important part in the infection process very few papers have been published regarding the movement of plant parasitic nematodes in different soil types in Australia. This study attempted to determine the movement of M. javanica and P. thornei, in different soil types under specific conditions, through pot culture studies using tomato as the host plant.

4.2 Materials and Methods

The culture of M. javanica inoculum was maintained on tomato, S. lycopersicum var. Grosse Lisse for the duration of the experiment. P. thornei cultures, maintained on carrot, were obtained from Dr Jennifer Cobon, DPI Queensland and used for the study. As they were maintained in culture, the population used for the experiment was assumed to be isogenic, thus eliminating variation which can arise from different populations.

Experiments were conducted using tomato cultivar Bite Size. Seeds of both Grosse Lisse and Bite Size varieties of tomato were purchased from Terranova Seeds Pty Limited NSW.

Three soil types representing a range of textures from heavy to light were selected from soil sample collections at UWS. These soils were subjected to detailed physical and chemical analysis. A summary of their major characteristics are presented in Table 4.1. The full soil analyses are presented in Appendix 1

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Table 4.1 Physical and chemical properties of sand, sandy clay loam and clay soils used in the nematode movement experiment

Soil 1 Soil 2 Soil 3 Texture Clay Sandy Clay Sand Class Loam % Sand 33.4 61.5 91.3 (SD*) (0.2) (0.2) (0) %Silt 6.0 15.9 0.7 (SD) (2.0) (0.2) (1.2) % Clay 60.6 22.6 8.0 (SD) (2.0) (0.2) (1.2) pH Water 6.5 5.0 6.1 (SD) (0.3) (0.2) (0.2) Electrical 0.2 283.0 0.1 conductivity (0.0) (1.2) (0.0) EC 1:5 dS/m (SD) % Total 0.9 3.5 0.6 organic C % Organic 1.5 6.0 1.0 matter % Total 0.05 0.3 0.0 organic N C:N ratio 17.6 11.0 18.0

*SD = Standard Deviation

Plastic pots of 1.5 L capacity (15 cm diam.) were filled with soil, up to 1 cm from the top edge, and then steam pasteurised at 60º C for 20 min. Sixty-three pots were set up for the experiment. These comprised 18 pots for each soil type, which were randomly allocated to the following treatments: no nematodes, nematode root counts 3 d after inoculation (DAI), 6 DAI and 10 DAI. Each treatment was replicated three times. One 7-day-old tomato seedling was transplanted into each pot about 2 cm from the edge. The pots were then placed on shelves in a temperature-controlled room (27±3º C) with artificial lighting (16 h dark; 8 h light) for 10 d. Watering was done daily at 09:00 (Eastern Summer Daylight Saving Time), to maintain the moisture of the soil just below field capacity. A saucer was placed underneath each pot for drainage of excess water. One week after transplanting, all plants except the control pots were inoculated with 300 nematodes each (either M. javanica or P. thornei) on the same day. Nematodes were applied as a 1 mL solution with a pipette, into a hole 5 mm deep, 10 cm away from the stem of each plant. In place of nematode inoculum, 1 mL of sterile tap water was added to the control plants. Plants were harvested at 3, 6 and

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10 DAI. Immediately after harvesting, roots were washed and processed for staining using acid fuschin according to the method of Byrd et al., (1983). The total number of nematodes in the root of each soil type was then counted under a light microscope at 100X magnification. Movement was measured by the rate of infection of nematodes in tomato roots.

4.2.1 Statistical Analysis

Data were analysed using two factorial analysis of variance (ANOVA) general linear SPSS® for WindowsTM Version 12 (SPSS Inc. 2003). The assumption of normal distribution was checked using P-P plot and homogeneity of variance checked using Levene’s test of equality of error variances (Levene, 1960).Treatment means were compared using Ryan’s Q test if the assumption of equality of variance was met or Dunnett’s T-test if the assumption was not met. Significance was accepted at the 0.05 level. Control pots were used to confirm absence of M. javanica or P. thornei in roots of potted tomato plants, in the treatment soils in the absence of nematode inoculation.

4.3 Results

4.3.1 Movement of Meloidogyne javanica to roots of Solanum lycopersicum in sand, sandy clay loam and clay soils There was significant difference between soils with regard to movement of Meloidogyne javanica to roots in sand, sandy clay loam and clay soil. Roots of plants grown in the sand and clay soils had significantly higher mean numbers of M. javanica at all time intervals than plants that had been grown in sandy clay loam soil

(F 2, 18 = 4.094, p = 0.034).

Three days after inoculation

At 3 DAI clay soil showed highest M. javanica infection of tomato roots, with a mean nematode count per plant of 22.3 ± 13.4 (SE) (7.4% of the initial inoculum) followed by sandy clay loam with a mean of 15.0 ± 5.9 (5%). The lowest number of

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M. javanica was recorded in roots of tomatoes grown in sand soil, with a mean nematode count of 9.3 ± 6.9 (3.11%) (Figure 4.1).

Mean number of Meloidogyne javanica in tomato roots in different soil types

20 3DAI

30 6 DAI6 20

90 80 70 60 50 40 10 DAI DAI 10 30 20 10 0 Sandy Sandy Clay Clay Loam

Figure 4.1 Mean number of Meloidogyne javanica observed in roots of tomato, Solanum lycopersicum cv. Bite Size, grown in sand, sandy clay loam and clay soil three, six and ten days after inoculation. (Standard Error of Mean for sand soil = 6.89, sandy clay loam = 5.86 and clay soil = 13.38).

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Six days after inoculation

At 6 DAI the highest number of M. javanica was found in roots of tomatoes grown in sand soil with a mean nematode count of 40.0 (13.3%) followed by clay soil with 29.0 (9.7%). The lowest mean number of M. javanica was recorded in roots of tomatoes grown in sandy clay loam soil with 12.0 (1.3%) (Figure 4.1).

Ten days after inoculation

At 10 DAI, the highest mean number of M. javanica was observed in roots of plants in sand soil with a mean nematode count of 55.7 (18.6%) followed by clay with 40.0 (13.3%). The lowest number of M. javanica (6.3 = 2.1%) was recorded in roots of tomatoes grown in the sandy clay loam soil (Figure 4.1).

4.3.2 Movement of Pratylenchus thornei to roots of Solanum lycopersicum in sand, sandy clay loam and clay soils

There was a significant difference between soils in the movement of P. thornei in sand, sandy clay loam and clay soils as measured by their presence in the tomato plants. Roots of plants grown in sand soil had significantly higher numbers of P. thornei than those grown in sandy clay loam or clay soil (F 2, 18 = 6.076, p=.0.01) (Figure 4.2).

Three days after inoculation

At 3 DAI, plants in the sand soil had the highest number of P. thornei in their roots, with a mean nematode count of 10.3 (3.4%), followed by plants in sandy clay loam with 6.3 (2.1%). Plants in clay soil had the least number of nematodes (0.7, 0.2%). (Figure 4.2).

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Mean number of Pratylenchus thornei in tomato roots in different soil types

10 3 DAI 3

20 18 16 14 12 10 6DAI 8 6 4 2 0

10 10 DAI 30

0 Sandy Sandy Clay Loam Clay

Figure 4.2 Mean numbers of Pratylenchus thornei observed in roots of tomato, Solanum lycopersicum cv. Bite Size, grown in sand, sandy clay loam and clay soil three, six and ten days after inoculation. (Standard Error of Mean for sand soil = 7.45, sandy clay loam = 0.0 and clay soil = 1.0).

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Six days after inoculation

At 6 DAI, the highest number of nematodes in tomato roots was recorded for those in the sand soil with a mean count of 10.7 (3.5%). Roots of tomatoes grown in the clay soil had an average of 1.0 (0.3%) nematodes while those grown in the sandy clay loam soil did not have any nematodes (Figure 4.2).

Ten days after inoculation

At 10 DAI the highest number of nematodes in roots was recorded in the sand soil with a mean count of 34.0 (11.3%). Roots of tomatoes grown in the clay soil had an average of 23.0 (7.6%) nematodes while those grown in the sandy clay loam soil did not have any nematodes (Figure 4.2).

4.4 Discussion

A comparison of the movement of M. javanica and P. thornei in sand, sandy clay loam and clay soils showed that the type of soil had significant effects on nematode movement and subsequent infection of tomato roots. The differences between treatments were consistent. The tomato variety used for the experiment was a good host for both M. javanica and. P thornei, as both nematode species successfully entered the host roots freely. M. javanica successfully induced giant cells and completed its life cycle within the roots (see Chapter 5).

Meloidogyne javanica

The percentage of the initial inoculum of M. javanica counted within roots of tomato plants grown in sand and clay soils steadily increased over 3 DAI, 6 DAI and 10 DAI (3.1, 13.3 and 18.6 % in sand soil and 7.4, 9.7 and 13.3 % in clay soil, respectively). These results are comparable to those reported by Hashmi et al (1994) who recorded 11 % penetration of tomato by Meloidogyne incognita after two weeks, in vitro. The largest number of M. javanica larvae counted 3 DAI in tomato grown in clay soil might have been due to the fast movement of the small sized larvae and also the availability of the necessary pore space and sufficient moisture. Additional factors

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which may have induced large number of nematodes to reach tomato grown in clay soil initially include the energy level of the nematode which decreased over time and the water retention capacity of clay soil. M. javanica has to utilise the energy stored in their body for all activities until they find a host as they do not feed until they have initiated feeding sites within the host root. Therefore faster and more active nematodes reached host roots first while the slower nematodes which had lesser energy reserves would have reached host roots only by the tenth day. Prot and Van Gundy (1981) observed that clay particles assisted in attracting M. javanica juveniles (J2) to the host plant over long distances by attracting and retaining root exudates. This, in addition to bacterial by-products in the root zone forms a concentration gradient, enabling nematodes to locate roots. Some similar action may have contributed to increased number of M. javanica observed in tomato plants grown in clay soil at 3 DAI. As moisture levels in pots were maintained at or near field capacity, it is possible that in clay soil the movement of nematodes may have been more in the horizontal direction rather than in sand soil which was probably in a more downwards vertical direction following watering after inoculation.

Observations at 6 and 10 DAI showed the largest numbers of M. javanica in roots of tomato plants grown in sand texture soil (13.33 and 18.56 %, respectively) compared to the two other soil types. Similar results have been reported previously (Verma and Jain, 1998; Desaeger and Rao, 2000) that M. javanica infection is more severe in sandy soils. Starr et al. (1993) found M. incognita more in coarse textured sandy soil with a pH of 7.5 to 8.5 and this supports the results of my study. However, contrary to their results, my study showed that more nematodes were able to travel through clay soil than sand in the initial three days after inoculation to reach roots of tomato plants.

Interestingly, in sandy clay loam soil the nematode numbers were consistently lower than both the sand and clay soils at 3, 6 and 10 DAI (5.00, 1.33 and 2.56 %, respectively). It was found, on analysis, that this soil had the lowest pH (5) and had, by far, the highest percentages of total organic carbon, total organic nitrogen, organic matter and the lowest C: N ratio (see Table 4.1). It also had the highest percentage of silt (15.7%). These characteristics may have affected the movement of the nematodes towards host roots and, thus, significantly reduced the level of root infection. Soil

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chemistry effects on the nematodes may have been due to the pH, organic matter and C:N ratio, while physical characteristics of the soil such as texture and particle size influenced nematode movement by affecting the pore size. Wallace (1960) reported a relationship between nematode movement, pore size, nematode diameter and water distribution.

Results obtained by Zasada and Tenuta (2008) show that organic amendments with urea and an alkaline stabilized biosolid suppressed the population of M. incognita in soil. They also reported that at high temperatures combined with soils of low water holding capacity, more ammonia was probably released resulting in nematode suppression. In my study the temperature was maintained at a constant 27 ± 3° C , and production of ammonia was not measured, so it could not be determined whether this was a factor in the low nematode count in tomato roots grown in sandy clay loam soils.

There may be other reasons for this low root population of M. javanica. It is known that Meloidogyne can change its sexual characteristics according to environmental conditions (Papadopoulou and Triantaphyllou, 1982; Triantaphyllou, 1993; Karssen, 2002), with unfavourable conditions resulting in more males. Sexual differentiation occurs in about the fourth larval stage and males can migrate out of roots (Abad et al, 2008). It is possible that a combination of temperature with other characteristics of the sandy clay loam soil resulting in more nematodes leaving the root, thus adversely affecting the number of nematodes within tomato roots.

Another possible reason for the low infection of tomato roots in sandy clay loam may be due to the presence of soil micro organisms which flourish in organic matter, including nematode-trapping fungi such as Arthrobotrys oligospora, Dactylaria psychrophila, as well as other species (Cooke, 1962; reviewed in Pramer, 1964). Jaffee and Muldoon (1995) reported that Monacrosporium spp. suppressed up to 92% of Meloidogyne javanica. In the present study, all soils were pasteurised prior to commencement of the experiment. This may have removed most soil biota, but some fungal structures may have survived in the soil and infected nematodes subsequently. The fact that pasteurisation removed most of the organisms may have also created

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conditions suitable for these types of nematode feeders to proliferate in the absence of their natural enemies and competitors.

Pratylenchus thornei

Similar to M. javanica, the percentage of initial inoculum of P. thornei counted within roots of tomato grown in sandy and clay soils steadily increased over 3 DAI, 6 DAI and 10 DAI (3.4, 3.6 and 11.3 % in sandy soil and 0.2, 0.3 and 7.7 % in clay soil, respectively). The highest number of P. thornei was consistently found roots of in tomato plants grown in sandy soil when compared to sandy clay loam and clay soils at 3, 6 and 10 DAI (Figure 4.2), showing that for P. thornei, sand was most suitable for moving towards and infecting tomato roots.

The relatively high percentage component of sand (91%) and the resulting large pore size could have supported movement of this nematode with its larger body diameter towards the host root, in the presence of sufficient moisture. Naganathan and Sivakumar (1975) reported that Pratylenchus thrives in sandy clay loam soils (even in the absence of plant hosts). This supports the results of the present study, which showed consistently low percentage counts of P. thornei in tomato roots grown in sandy clay loam soil at 3, 6 and 10 DAI (0.2, 0.0 and 0.0 %, respectively). As P. thornei is a migratory ecto-endo parasite, it can feed on any host root and survive easily in soil. The low population of P. thornei in plant roots, despite the presence of visible root damage, may be due to their feeding on the host roots and then exiting after feeding. In well-watered plants, Pratylenchus may be encouraged to migrate outside the roots, compared to water-stressed conditions. In my experiment, the tomato plants received adequate water. Another possible reason for low nematode population in roots of tomato plants grown in the sandy clay loam soil, is that it had, by far, the highest level of organic matter of all the soils tested. This may have lead to higher nematode-suppressing properties associated with by-products of organic matter breakdown such as ammonia, and presence of antagonistic biota, as has been previously described for M. javanica plays some similar role here. However, I did not assess survival of nematodes in the soils during the experiment.

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Overall, results of this study indicated that many factors influence the movement of M. javanica and P. thornei towards tomato roots:

(a) soil characteristics such as texture, and chemical composition, moisture (b) diameter of the nematode body and its capacity for sensory perception to detect possible signals from the host root.

The soils used in this experiment were typical of Australian soils, many of which are naturally low in nutrients and organic matter. However, in farming situations, there may be high variation in soil physical and chemical characteristics, resulting from the initial soil formation as well as agricultural practices, such as tillage, fertiliser use and incorporation of organic matter. Based on my results, these factors may significantly affect nematode movement towards roots of potential host plants.

Thus, organic soil amendments could augment other strategies, such as soil solarisation and/or crop rotation with non hosts, as part of integrated nematode management. Results of this study support the view that incorporation of organic matter into soils may reduce the severity of plant parasitic nematode infection, even in soils that would otherwise be conducive to crop damage by nematodes. If successful, this could result in a reduction in environmental pollution by the reduction in the amount of toxic pesticide chemicals, increased crop yield, and, ultimately, increased farmers’ incomes.

Given the importance of root parasitic nematodes and wide range of soil types in world crop production systems, this study indicates that this relationship needs to be explored further in both laboratory and field investigations.

4.5 Conclusion

Sandy clay loam soil was least suitable for both species of nematodes for their location and infection of tomato roots. This soil had a pH of 5, the highest percentages of total organic carbon, total organic nitrogen, organic matter and the lowest C : N ratio, as well as having the highest percentage of silt. I conclude that some or all of these characteristics played major role(s) in reducing the number of M.

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javanica and P. thornei able to move through the soil and infect tomato roots, in the presence of adequate soil moisture. Thus, soil texture was not the primary factor.

Size of the infective juveniles, particularly body diameter, as well as their genetics, virulence and energy reserves can influence movement of nematodes through soil. However, it was not clear to what extent they influenced nematode movement towards tomato roots in this study. Presence of other microorganisms and chemical by-products, associated with this soil’s high organic matter content, may have also played a part in reducing the number of nematodes reaching the host plant roots.

This study demonstrates the importance of chemical characteristics, as well as soil texture, in influencing nematode movement towards host roots. It further suggests that modifying soil by using organic amendments may play an important role in integrated management of plant parasitic nematodes.

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CHAPTER 5

Temporal study of the early infection of tomato, Solanum lycopersicum, by Meloidogyne javanica and Pratylenchus thornei

5.1 Introduction

When nematodes infect host plants they breach several defense mechanisms that the host has developed in its evolutionary history to ward off harmful invaders (Reviewed by Levin [1976] and more recently by Trudgill [1991]). Over the years several detailed light microscope and live video imaging studies have been carried out to elucidate the host-parasite interactions between nematodes.

Three days after infection with Meloidogyne incognita, seedlings of Impatiens balsamina examined in light microscopic studies showed early stages of giant cell formation. These giant cells were up to six times the length of the neighbouring cells. By day 6 giant cells had expanded considerably, and by day 10 they had reached up to 600 m. A parallel study by transmission electron microscopy revealed wall ingrowths in giant cells close to xylem and sieve elements and appeared to overlie walls of non-conducting parenchyma cells near them. Between days 3 and 6 after initiation the pit fields between giant cells and the cells outside them are lost, but a specific pattern of pit fields is formed between adjacent giant cells. By day 17 nematodes were many times larger than the giant cells. By day 30, there were distinct signs of wall degeneration in giant cell walls (Jones and Dropkin, 1975).

Video-enhanced contrast light microscopy and time lapse studies (Wyss 1992) of the parasitic behaviour of second-stage juveniles (J2) of Meloidogyne incognita in roots of Arabidopsis thaliana showed that J2 invaded the root primarily in the region of elongation, close to the meristematic zone. Epidermal and subepidermal cells were destroyed during invasion, while intercellular invasion between epidermal cells was less frequent. After invasion, the J2 within the root were always oriented in the direction of the root tip and migrated towards it between cortical and meristematic cells without causing any damage. Wyss observed occasional attempts at intracellular

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migration through cortical cells, but these were not successful. When the J2 reached the apex of the root, they turned round and migrated away from the root tip between the cells towards the differentiating vascular cylinder. Within the differentiated cylinder migration eventually stopped and giant cell induction was initiated. After a period of induction, the J2 became surrounded by young multinucleate giant cells within about 24 h. A characteristic behaviour pattern, composed of continuous head and stylet movements, interspersed by periods of stylet tip protrusion and metacorpal bulb pumping, was maintained throughout all phases of parasitism.

On cabbage seedlings, P. penetrans congregated around roots and started feeding 3 h after inoculation. Faint yellow lesions appeared on roots as early as 24 - 36 h after inoculation. Cell walls at the feeding sites were thicker than normal. The nematode moved intra- and inter- cellularly. Initially, for up to 30 d after infection, the endodermis provided a barrier to Pratylenchus, but after this the stele was invaded as well (Acedo and Rohde, 1970). Kurppa and Vrain (1985) observed Pratylenchus penetrans behaviour infecting strawberry roots under a stereo microscope. They observed that most nematodes moved to the root within 3 h. They explored the root surface with their stylets and pierced holes in a cell for entry, mostly avoiding root tips or the elongation zone. In some cases a line of holes were made on root hairs, and then a hole was torn by pressing their head and waving it vigorously. The cytoplasm of these cells circulated vigorously during and after nematode penetration, then the cells collapsed rapidly. Once the stylet was inside a cell, a globular secretion passed from the stylet tip into the cell, and the movement of the cytoplasm became more agitated. Nematodes moved from the epidermal cells to the cortical cells and then longitudinally along the length of the root, each time making a row of holes and pushing their head into the adjacent cells.

Zunke (1990) used high resolution video-enhanced contrast microscopy to study the behaviour of P. penetrans on root hairs of four different plant species (rape, oil radish, tobacco and potato). The behaviour was separated into phases of probing, cell penetration by the stylet, salivation and food ingestion (for brief and extended periods). After cell penetration a small “salivation zone” was formed around the stylet tip. Infected root hairs showed increased rate of cytoplasmic streaming and

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gradual hypertrophy of the nucleus. Defaecation occurred every 2 - 4 min during extended feeding periods.

As mentioned above, many studies have been previously conducted on a number of aspects of the plant-nematode infection process. However, there are several aspects of these interaction mechanisms still to be fully investigated, such as early pathogenesis. In addition, there have not been any comparative studies between Meloidogyne and Pratylenchus, with regard to the infection process. This study was derived from the larger nematode movement study in different soil texture classes, described in Chapter 4. It aimed to follow, through a temporal sequence, the early infection process for both M. javanica and P. thornei.

5.2 Materials and Methods

The M. javanica culture was maintained on tomato, S. lycopersicum cv Grosse Lisse, plants for the experiment using methodology previously described in Chapter 2. P. thornei inoculum for the study was extracted, as necessary, from stored cultures originally obtained from Dr Jennifer Cobon, DPI Queensland and maintained on carrot (Chapter 2).

The methodology for the study is essentially that described in Chapter 4. The host plants were tomato cultivar Bite Size. However, for this temporal study, only plants grown in sandy soil were selected. This was to avoid confounding of the results arising from interactions of different soil texture classes, and because it gave sufficient clear images. Plants from the heavy clay were rejected because it was difficult to wash roots adequately to enable high quality images from intact roots; plants from sandy loam recorded, by far, the lowest number of nematodes in roots. Plants were harvested on the 3rd, 6th, 10th and 18th day after inoculation with nematodes. Immediately after harvesting, roots were washed and processed for staining using acid fuchsin and processed according to the method of Byrd et al. (1983). Under a stereomicroscope (Leica MZ12), the roots were placed on to microscope slides on a drop of glycerol and coverslip was gently placed onto the root excluding any air bubbles. Samples were usually examined as close to preparation as possible. In some cases when this was not possible, samples were stored in fresh

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glycerol until required. These slides were then placed under an Olympus BX60 compound microscope and images at various magnifications were collected under transmitted light on a Jenoptik ProgRes C14 digital camera, using the software Image Pro Plus Version 7 (Media Cybernetics Inc., Bethesda, MD, USA).

5 3 Results

The following information relates to observation of images, from the different sample dates. Typical images are presented to illustrate different nematode stage(s) and their sequential development, as well as nematode migration within plant roots and subsequent tissue damage. In several cases, images from the confocal studies using Arabidopsis (Chapter 2) have been included, for comparative purposes.

5.3.1 Early infection of tomato, Solanum lycopersicum, root by Meloidogyne javanica

Observations 3 days after inoculation Observation of M. javanica- inoculated roots as early as 3 DAI showed several J2 migrating within the root (Figure 5.01, Figure 5.02). Within the cortex, several nematodes which had reached the root tip turned into the vascular bundle (tn) and became oriented towards the central vascular cylinder. J2 already within the central cylinder were oriented parallel to one another and to the long axis of the root and had started migrating towards the region of differentiation to find a suitable feeding site (nb). During the initial migration of J2 towards the root tip there was negligible damage to the host tissue (Figure 5.03). One J2 was observed with its head region (nh) positioned between two cell walls (cw).

In one root several invading J2 were seen breaching the central vascular cylinder directly (Figure 5.04), rather than following the initial normal path towards the tip, to continue their migration towards a potential feeding site. In the root cortex, the migratory path taken by other nematodes was markedly visible. During their migration they caused damage to several xylem vessels (dx).

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Figure 5.01 Light micrograph of tomato, Solanum lycopersicum, root shows Meloidogyne javanica second stage juveniles (J2) migrating within 3 days after inoculation.

Root tip is swollen due to simultaneous infection by a large number of nematodes (st); one nematode which has reached the root tip is turning into the vascular bundle (tn); within the differentiating region, nematodes are aligned parallel to one another and migrate to the differentiating region to find a suitable feeding site (nb1). Sample was stained with acid fuschin. Observed under 20X magnification.

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Figure 5.02 Higher magnification of root tip in Figure 5.01.

Damaged, swollen root tissue (st); J2 turning into the vascular bundle (tn); J2 migrating within the vascular bundle towards a potential feeding site in the differentiating region (mn). Samples were stained with acid fuschin. Observed under 40X magnification.

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Figure 5.03 Transmitted light micrograph of a Meloidogyne javanica J2 migrating within a tomato (S. lycopersicum) root.

J2 is positioned with its head (nh) between two cell walls (cw). Observed under 400X magnification.

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Figure 5.04 Transmitted light micrograph of a tomato, Solanum lycopersicum, root shows several invading Meloidogyne javanica J2.

The J2 have breached the vascular bundle (nv), directly damaging the xylem vessels (dx). Observed under 100X magnification.

Observations 6 days after inoculation

Although inoculation of tomato was carried out at only one time, nematodes entered the roots at different times. The next image (Figure 5.05) was taken at 6 DAI and the nematode seen was still migrating to find a suitable feeding site. A fine stylet could be clearly seen (st2). A discontinuous xylem vessel was close to the head of the J2, possibly disrupted during infection (dx).

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Figure 5.05 Transmitted light micrograph of a tomato, Solanum lycopersicum, root shows Meloidogyne javanica J2 migrating within the vascular bundle.

The head of the J2 (nh) is situated close to a discontinuous xylem vessel (dx). Stylet of the nematode (st2) is clearly visible along with other xylem vessels (x). Observed under 400X magnification.

Observations taken at 6 DAI showed late stage J2 within roots. The nematode (Figure 5.06) had started its transformation from the J2 to J3 stage. A curled tail possibly indicated the initiation of moulting (nt). The J2 was situated with its head (nh) in the vascular bundle and the curled tail (nt) within the cortex. A newly forming lateral root developed close to the head of the J2, and new wound xylem (xf) was formed around the feeding site; undisrupted xylem also was seen (x) away from the infection site.

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Figure 5.06 Transmitted light micrograph of a tomato, Solanum lycopersicum, root shows a Meloidogyne javanica juvenile in a stage between a late J2 and an early J3.

The J2 is situated with its head (nh) in the vascular bundle and the curled tail (nt) within the cortex; close to the head of the J2, a newly forming lateral root (lr) can be seen; Xylem vessels develop around the feeding site (xf); undisrupted xylem is also seen (x). Observed under 100X magnification.

Some nematodes were observed within this time period in the process of moulting from the infective J2 to the J3 stage (Figure 5.07).These probably were nematodes which entered the root earlier and therefore had an earlier start to their development. The head of the nematode (nh) was indistinct as it was embedded deep within root tissue. In this image, outline of the nematode intestine could be seen (ni). The tail region (nt) of the nematode body was rounded as though it had constricted towards the head region. Nematodes in J3 stage had a thicker body diameter than J2. The old,

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empty cuticle was still visible and was clearly stained near the tail region (nt). Old cuticle seemed to be peeling away from the nematode body (cb).

Similar observations were made in transmitted light images during cytoskeletal studies of whole roots of Arabidopsis using confocal microscopy (Figure 5.08). The tail of the J2 in this image made two distinct impressions in root tissue when observed at two different depths from the root surface. In this image, arrows show an imprint of the tail made in root tissue during the nematode movement and arrowheads show the actual position of the tail.

Similar to the curled tail observed earlier in Figure 5.06 at a stage where the nematode is undergoing a change in body shape, these positions of the nematode tail within the tomato root may signify the beginning of the moult from J2 to J3. A and B are confocal optical sections taken 26 µm and 33 µm from the root surface. Bar = 30 µm.

Observations 10 days after inoculation Several late J3 and J4 stage juveniles were observed in roots ten days after inoculation. The J4 in one case (Figure 5.09) was located with the head in the vascular tissue and the body towards the cortex (c). A distinct neck (nn) could be delineated between the head (nh) and the thick body (nb). The nematode head was located amidst several large giant cells (gc).

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Figure 5.07 Light micrograph of a tomato, Solanum lycopersicum, root shows a Meloidogyne javanica J2 moulting into a J3.

The head of the nematode (nh) is indistinct as it is embedded deep within root tissue; empty cuticle was seen in the tail region (ct) along with the shedding cuticle of the body (cb); outline of the nematode intestine was clearly visible (ni); the tail region (nt) of the nematode body was rounded as though it had contracted towards the head region. Observed under 100X magnification.

Figure 5.08 Transmitted light confocal image of Arabidopsis thaliana root showing early stage of Meloidogyne javanica J2 moulting.

Arrowheads show the actual position of the nematode tail and arrows point to imprints made in root tissue during the nematode movement. (A and B are confocal optical sections taken 26 µm and 33 µm from the root surface). Bar=30 m.

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Figure 5.09 Light micrograph of a tomato, Solanum lycopersicum, root showing a late stage Meloidogyne javanica J3 within root tissue.

Head of the J3 (nh) is embedded within the vascular tissue (vb); a clearly discernible neck (nn); the body of the J3 is situated outside the vascular bundle, in the cortex (c). Observed under 100X magnification.

Figure 5.10 Transmitted light micrograph of a Meloidogyne javanica J3 within a tomato, Solanum lycopersicum, root.

A lateral root is forming close to the feeding site (lr); vascular bundle (vb) and the cortex (c); nematode head (nh); nematode neck (nn). The vascular bundle shows a slight swelling in the region of feeding site formation; nematode head is situated amidst the giant cells (gc); nematode neck; distinct median bulb (mb) of the nematode which acts as a pump during feeding; Images A and B. Observed under 100X and 200X magnification, respectively.

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Emergence of a lateral root in the cortex close to the feeding site was observed (Figure 5.10, A). This was similar to observations made 6 DAI in Figure 5.06. Observation at a higher magnification (Figure 5.10, B) revealed distinct large giant cells (gc) around the head region of the nematode. A well developed median bulb (mb) was observed within the nematode body (nb) just below the distinct neck (nn) region. The nematode had its body mostly contained within the vascular bundle (vb). The vascular bundle showed a slight swelling in the region of feeding site formation.

Observations 18 days after inoculation M. javanica was observed moulting from late J3 to the J4 stage about 18 days after inoculation (Figure 5.11). A feeding site (gc) was located within the vascular bundle (vb); the nematode was situated with its head embedded within the vascular region and the body (nb) in the cortex (c); a neck region could be distinguished (nn); the nematode body (nb) had started to resemble the typical spherical adult female form; the tail region (nt) had constricted even more towards the anterior region of the body and left the shadow-like image of an empty cuticle (ct) behind.

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Figure 5.11 Transmitted light image of a tomato, Solanum lycopersicum, root infected by Meloidogyne javanica.

The nematode is moulting from late J3 to the J4 stage; A distinct feeding site (gc) is located within the vascular bundle (vb); the nematode is located with its head embedded in the vascular region and the body (nb) in the cortex (c); a neck region can be distinguished (nn); the nematode body has started to resemble the typical spherical adult female form; the tail region (nt) has constricted even more towards the anterior region of the body and has left an empty cuticle (ct) behind. Observed under 100X magnification.

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Figure 5.12 A tomato, Solanum lycopersicum, root with an adult Meloidogyne javanica within.

The body of the nematode (nb) is situated in the cortex; the nematode body is completely surrounded by cortical tissue (cg) forming the gall; cells in the gall are of a different, irregular shape compared to normal cortical cells (c); the vascular bundle (vb) encloses the feeding site of M. javanica (gc). Observed under 40X magnification.

Some nematodes observed within roots 18 DAI were in mature adult stage (Figure 5.12), with the body (nb) situated in the cortex and completely surrounded by cortical tissue (cg) forming the gall. Cells in the gall were of a different, irregular shape compared to normal cortical cells (c). The vascular bundle (vb) enclosed the feeding site of M. javanica (gc).

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Figure 5.13 An adult Meloidogyne javanica within a tomato, Solanum lycopersicum, root.

The adult nematode body (nb) is partially surrounded by cortical tissue (c); an enlargement in the vascular bundle (vb) encloses the feeding site composed of giant cells (gc); the disrupted cortical region (dc) is probably where an egg mass was dislodged earlier during sample processing. Observed under 40X magnification.

Adult M. javanica lay their eggs to the outer surface of the root. Although several of these adult nematodes had egg masses attached, the egg masses became detached (Figure 5.13) during the sample preparation process revealing damaged tissue on the root surface (dc).

5.3.2 Early infection of tomato, Solanum lycopersicum, root by Pratylenchus thornei

As in the case of Meloidogyne, inoculation of Pratylenchus was done simultaneously on tomato, although P. thornei entry into tomato roots was observed throughout the

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investigation period. While this also occurred in M. javanica, it was much more common in Pratylenchus. This, together with the similar morphology between stages of Pratylenchus, made it more difficult to develop a similar timed sequence of events for this species.

Observations 3 days after inoculation

Observations taken at 3 DAI showed P. thornei migrating within roots through the cortex. The head of a nematode was pressed firmly against an end (cross) wall (cw) (Figure.5. 14). Some substance (possibly secretory material) was observed extruding (sm) from the nematode (n) into the plant cell.

Observations 6 days after inoculation

In almost all tomato roots inoculated with P. thornei, many necrotic yellowish brown lesions were observed in root tissue, and the path in the cortex along which the nematode migrated could be easily distinguished in some roots by the damage caused to the cells. Roots observed at 6 DAI with P. thornei showed several damaged epidermal cells on the surface which were probably the entry points of the nematodes (ne) into the root tissue (Figure 5.15).

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Figure 5.14 Tomato, Solanum lycopersicum, root infected by Pratylenchus thornei.

The nematode (n) is breaching the end wall of a cell (cw)in the cortex (c); it appears as though some material (sm) is extruding from the head region of the nematode into the cell cytoplasm. Observed under 400X magnification.

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Figure 5.15 A tomato, Solanum lycopersicum, root observed six days after inoculation with Pratylenchus thornei.

Image shows several damaged epidermal cells on the root surface which may be the entry points of the nematodes (ne) into the root tissue.

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Figure 5.16 Pratylenchus thornei infecting a tomato, Solanum lycopersicum, root.

Damaged epidermal and cortical region of the infected root (dc); a migrating nematode (n); damage localised to a single cell (ce), probably entry points of the nematode into the root tissue; highly stained unidentified structures (st1), probably excretory matter from P. thornei. Observed under 100X magnification.

Epidermal and cortical tissues of tomato root were damaged during the infection by Pratylenchus thornei (Figure 5.16). In some cases damage was localised to a single cell (ce), probably entry points of the nematode into the root tissue while in some others a group of cells over a significant area was affected, showing a yellowish brown discolouration (dc). Some highly stained unidentified oval structures (st), probably excretory matter, from P. thornei were observed within root tissue.

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Figure 5.17 Surface tissue of tomato, Solanum lycopersicum, root damaged due to infection by Pratylenchus thornei.

The lesions caused by P. thornei have soil firmly attached which was not dislodged even after the staining process. Observed under 40X magnification.

Some roots had soil particles attached to the surface lesions which had not been displaced even after the whole staining procedure (Figure 5.17).

Observations 10 days after inoculation

As described in an earlier paragraph, P. thornei was seen to enter the roots all through out the duration of the experiment.

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Figure 5.18 Pratylenchus thornei infecting a tomato, Solanum lycopersicum, root.

Three P. thornei nematodes can be seen in this image. One is seen in the process of entering the roots (ne), while two are seen migrating within cortical tissue (nm) outside the vascular bundle (vb). Observed under 100X magnification.

Observations taken at 10 DAI (Figure 5.18) showed a nematode in the process of entering a root (ne), with part of its body within and part outside the root. In the same image two P. thornei could be seen migrating (nm) within the cortex. Nematodes were not observed within the vascular bundle.

Although Pratylenchus moved fairly in a straight line through tomato root, it was sometimes observed to be coiled in unusual positions in a single cell within the damaged tissue (Figure 5.19).

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Figure 5.19 Pratylenchus thornei within tomato, Solanum lycopersicum, roots.

Single nematodes (n) were observed in several cases curled up within a single cell enclosed on all sides by cell walls (cw). Observed under 400X magnification. Images A, B, and C are of tomato at 6 DAI and image D of tomato at 10 DAI.

5.4 Discussion

In order to elucidate the host-parasite interaction between plants and parasitic nematodes it is essential to have a good understanding of the early infection process. Several studies have been made earlier with Meloidogyne and Pratylenchus (Chapter 1). In this study an attempt was made to compare the sequential early infection processes of two plant parasitic nematodes, Meloidogyne javanica and Pratylenchus thornei on tomato, S. lycopersicum cv. Bite Size. It was easier to study the sequential pattern of M. javanica infection process based on the different

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morphology in its different life stages, whereas in the case of P. thornei a sequential study was more based on observations over time as the morphology of the different life stages were quite similar.

Results showed significant differences in the early stages of infection. Both M. javanica and P. thornei seemed to prefer some roots on the same plant to other roots for infection. Large numbers of nematodes were found in a few roots while some other roots had very few or no nematodes. Entry and migration of M. javanica showed minimum disruption or discolouration to root tissue initially, where as P. thornei entry and migration resulted in localised and widespread tissue destruction and discolouration. Within root tissue, migration of Meloidogyne was mainly between two adjacent cell walls. This supports Endo and Wergin’s (1973) findings that cells were separated along the middle lamella during migration. Cell wall degrading enzymes such as pectate lyase have earlier been isolated from the oesophageal glands and the cuticle surface of Meloidogyne (Doyle and Lambert, 2003). The median bulb acts as a pump during feeding, and was well developed in M. javanica.

Within the root, the head region of Pratylenchus was almost always observed pressed to, or oriented towards the cross walls of cortical cells. Secretions were noted close to the lip region of the nematode in the infected host cell. A similar observation was made by Zunke (1990) who reported a “salivation zone” in root hairs, close to the stylet tip of P. penetrans, which probably included accumulated host cytoplasm and nematode salivary secretions. In many images, a single Pratylenchus was seen to be enclosed, curled up, within a single cortical cell. Similar observations have been made in previous studies on various host plants by Zunke (1990) who reported that feeding and migration of P. penetrans were interrupted by rest phases when a nematode became characteristically coiled inside a cell. In most cases in my study, at least one cross wall of the cell surrounding the nematode was damaged, probably by prior nematode migration.

Initial migration of Meloidogyne was mainly through the cortex, although the final destination was always the vascular system of the host. Similar observations have been made by several earlier studies (Chapter 1). However, in my study it was

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observed that, in some cases, Meloidogyne directly breached and entered the vascular system in their quest for the ideal feeding site. This is consistent with the extensive disruption to the xylem observed to be caused by M. javanica on A. thaliana roots in transmission light images from cytoskeletal studies (Chapter 2, Figure 2.18).

Pratylenchus left the vascular system largely untouched and intact. Similar observations were made by Castillo et al. (1998) who reported that Pratylenchus migrated through epidermal and cortical cells by breaking down cell walls along the path of migration. However they did not mention whether it was the end walls or the lateral walls that the nematodes broke down. My study showed that Pratylenchus mostly targets the end walls of cells in the cortex. The reason for this is unknown. End walls result from cell plate formation during cell division. It is quite difficult to study the end walls of plant cells mainly due to their small thinness and small size. Therefore, only a limited number of studies have been conducted on end walls of cells [David Collings (2008), Personal Communication]. However, it is an accepted fact that end walls are physiologically and structurally different from the lateral walls (Wasteneys and Collings, 2006) because they have very different growth patterns and functions. Microtubules in lateral walls are arranged transversely while those at end walls are arranged radially. Actin microfilaments in end walls are found in a random arrangement. Pratylenchus may respond to some particular physical or chemical signals emitted by these cell walls.

Within Pratylenchus- infected roots an oval mass, staining highly with acid fuchsin, was noted in several cases, probably excretory material from the nematode. Acid fuchsin stain is specific to animal muscle tissue, connective tissue and collagen (Puchtler and Sweat, 1964). Zunke (1990) observed defecation by P. penetrans every 2-4 min during feeding. However, these oval unidentified structures were not observed in Meloidogyne-infected roots.

The juvenile stages of Meloidogyne moulted within root tissue, continuing the life cycle, and the outline of the cuticle being shed was clearly visible, in a few instances. This contrasts with Bird’s (1959) observation that some nematodes do not shed their old cuticle. Similar observations on moulting were not made in Pratylenchus- infected roots.

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The acid fuchsin stain from nematodes in samples stored in glycerol was found to spread into adjacent root tissue, with time. Nematode tissue shrinkage in stored samples was also noted, more so in Meloidogyne than in Pratylenchus.

5.5 Conclusion

A temporal study on early stages of infection by, and migration and development of, Meloidogyne javanica and Pratylenchus thornei on tomato, Solanum lycopersicum, revealed significant differences between the species. These were primarily in the way tissue in the plant roots was damaged. P. thornei caused extensive damage and lesions to the root surface and cortex while migration. M. javanica, however, appeared to cause minimal disturbance to the arrangement of cells during its early migration, although in the later stages it induced its feeding site and caused other structural changes in the root tissue, remaining undetected by the plant as an enemy for the whole of its life cycle. This difference in damage is likely to be reflected in the subsequent growth and development of infected host plants.

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CHAPTER 6 General discussion

This chapter discusses the major outcomes of my work that has been presented in the previous chapters, and the implications and potential applications emanating from my results.

6.1 Major findings / Outcomes

Plant parasitic nematodes cause damage to the plants they infect. There are several aspects of these interactions which are still unknown. My studies attempted to understand three different aspects of these interactions, namely • changes occurring in the plant root cytoskeleton due to nematode infection, . • effects of nematode infection of roots on the growth and development of tomato seedlings • movement of nematodes towards plant roots in different soil texture types

The nematodes chosen for this study were Meloidogyne javanica (root-knot nematode) and Pratylenchus thornei (lesion nematode), which had contrasting biology and modes of infection.

In Chapter 2, I reported on investigations of changes in the cytoskeleton of A. thaliana root cells following infection by these two nematode species. For these investigations, I used three techniques. The first was whole root immunolabelling using a methodology developed by Collings and Wasteneys (2005) for observing actin and microtubules simultaneously; the second was dual immunolabelling of BMM resin-embedded sections of infected tissue using a modified method developed by Baskin et al (1992); and the third utilised GFP-hTalin-transformed A. thaliana.

Whole root dual immunolabelling had not been previously used to study effects of nematode infection on the plant cytoskeleton. Using this technique, I found that actin microfilaments reacted differently to microtubules in surface wounds caused by

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nematode infection on roots. Initial responses of the plant to both M. javanica and P. thornei were similar. Both species of nematodes were observed to destroy the outer epidermal cell wall of the A. thaliana root during entry. In wounds confined to a single cell a round or disc-shaped plug, positively labelled for actin, was formed on the root surface and lined the inside of the wounded cell. Cortical microtubules in the affected cells, on the other hand, held their position, probably providing support to the remaining parts of the damaged cell wall. These observations were made within 3 h of inoculation. This is the first time that such cytoskeletal reactions have been reported following nematode attack on the surface of Arabidopsis roots.

Increased fluorescence was observed in all infected tissues, mainly in areas of nematode entry and feeding site formation, with all three techniques used. De Almeida Engler et al. (2004) suggested that such response is due to increases in tubulin and actin concentrations in infected cells. However, my study suggests that the increase in fluorescence may be due to some nematode-induced changes occurring in infected cell walls, resulting in increased permeability to primary or secondary antibodies used to target microtubules and actin microfilaments and, as a result, increased labelling of these proteins. This increased cytoskeletal labelling observed at the initial entry and final feeding site formed was, for the most part, noticeably absent in plant cells along the path of migration of nematodes. However, in the few cells in which fluorescence was observed, the microtubules did not have the normal taut arrangement; rather, they were seen to be in a wavy pattern. This is the first time that such an observation has been made. This may be a result of disruption or displacement of the host cells caused by the nematode forcing its way through the root tip. In addition, there may have been some degree of tissue rearrangement in the host in relation to the nematode during fixation or other microscopic procedures (David Collings, Personal Communication). Cell wall- loosening enzymes, reportedly produced by M. javanica during migration to separate the cells along its path, may also play a role in causing this effect. The different enzymes reported include expansins (Gal et al., 2006); pectin acetyl esterase (Vercauteren et al., 2002) and many others such as endoglucanases and xylanases (reviewed in Punja et al., 2008).

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I found that actin fixation was very difficult and, in cases where fixation was not adequate, a diffuse fluorescence was observed in place of well-defined microfilaments. De Almeida Engler et al. (2004) also observed diffuse fluorescence in infected tissue but they postulated that this was due to depolymerisation of actin microfilaments.

In the feeding sites, transverse cortical microtubules, several abnormal spindles and phragmoplasts were observed within the giant cells, but pre-prophase bands were not observed. These observations support those made by De Almeida Engler (2004). Other structures, such as the transfer cells of the Arabidopsis and other plants, have been previously reported to occur naturally (more details in Chapter 2). Thus, M. javanica has the ability to harness and modify this ability to form structures through as yet unknown biological pathways to produce a suitable environment for its own growth, development and reproduction.

The optical properties of cell walls in infected cells were conspicuously different to those of uninfected cells in transmission light images. This may be due to changes in the cell wall structure or chemistry, as a response to nematode infection.

Well defined and transversely arranged cortical microtubules were observed, using whole root dual immunolabelling, in the early stages of feeding site formation of M. javanica. Using BMM sectioning and dual labelling of infected tissue, microtubules and actin microfilaments were also observed. Actin microfilaments were also observed in gall tissue in the GFP-hTalin transformed Arabidopsis. Thus, the presence of well-defined microtubules and actin microfilaments in nematode- infected root tissue was confirmed using all three (different) techniques. During the later stages of infection this well-defined structure of microtubules seemed to deteriorate.

Cortical microtubules in giant cells were transversely arranged. This arrangement is similar to the transverse arrangement of microtubules observed in actively dividing meristematic tissues of the root. However, cell division was not achieved. The surrounding normal cells had the typical oblique arrangement of differentiated cells. This arrangement of microtubules has not been reported previously.

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Whole root immunolabelling showed that giant cells induced by M. javanica seemed to have lost their growth polarity and anisotropy. The giant cells resembled a spindle in shape, with an enlarged middle section and tapering towards both ends. This indicated a difference exists between the side walls and end walls of cells, and that the factors necessary for the expansion of the middle section of the cell may not be the same as those needed for the expansion of the end walls. This observation has also not been previously reported in nematode-infected tissues. However, my results are supported by Collings and Wasteneys (2005) who reported that cross walls of anisotropically expanding cells have fundamentally distinct properties to cell walls parallel to the longitudinal axis, confirming that side walls have different properties and characteristics compared to end walls.

Severe disruption was noted in the cell files and also in the quiescent centre when nematodes invaded the root tip in high numbers. In root tips, alternating cell files of trichoblasts and atrichoblasts were distinguishable. Severe reduction in the rate of growth in infected roots was indicated by the presence of well-defined xylem vessels and epidermal root hairs just behind the root tips. However, even though xylem vessels were disrupted extensively, they regenerated in a relatively short time (approximately 3 h). These observations have not been reported earlier. Disruption of the xylem may also be explained by the failure of differentiation of progenitor cells into xylem cells or non-elongating cells developing as wound xylem (Mike Jones, Personal Communication).

6.1.1 Comparison between the effects of Meloidogyne javanica and Pratylenchus thornei on Arabidopsis thaliana and Solanum lycopersicum

I conclude that Arabidopsis is a good host plant for cytoskeletal studies using M. javanica. However, it is not a suitable host for studying the infection process of P. thornei, as these nematodes were not observed within roots. Arabidopsis roots have a small diameter which is probably unsuitable for entry of large nematodes such as P. thornei. Tomato, with a much larger root diameter, would have been a more suitable plant host for studying these interactions with P. thornei.

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Whole root dual immunolabelling, dual immunolabelling of BMM resin-embedded sections and GFP-transformed A. thaliana were all effective in studying the effects on host cytoskeleton in the early stages of infection. However, each had its own advantages and disadvantages.

Whole root dual immunolabelling gave a good understanding of the 3D structure of both the nematode and the infected tissue. As mentioned previously, A. thaliana responded similarly to initial infection by both nematode species. The outer epidermal cell wall was destroyed during entry and a distinct wound plug, labelled positively for actin, was observed on the root surface and lining the wound. The actin microfilaments may have reacted to the breach of the cell wall by oozing out and accumulating on the surface of the wound. On the other hand, by holding their transverse position in the affected cells, cortical microtubules probably provided support to the remaining parts of the damaged cell wall. My microscopic observations with transmitted light of infected tomato roots supported the results I obtained in Arabidopsis because, in most cases, entry wounds were restricted to single cells in tomato.

I was unable to determine whether P. thornei produced the wavy microtubules I observed in Arabidopsis cells lining the migratory path of M. javanica, because no P. thornei were present in the Arabidopsis roots.

The initiation and development of giant cells and galls, associated with M. javanica infection, appeared to follow a similar path in A. thaliana and in tomato roots. In both hosts, cells around the nematode head region were transformed into giant cells (i.e. cell division without cytokinesis), while cells around the posterior body of the nematode were smaller and more numerous.

For M. javanica, life stages could easily be distinguished. However, this differentiation was difficult for P. thornei, due to similarities in the morphology of its different stages (Chapter 5). However, it was interesting to note that while Meloidogyne used naturally-occurring plant physiological processes to manipulate the host and remain undetected, Pratylenchus had a simpler and more basic method for infection; namely, entering, feeding and then leaving the root, causing substantial

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tissue destruction. Meloidogyne mainly affected the vascular system, whereas Pratylenchus preferred the cortical tissue for its infection.

6.1.2 Understanding the effects of nematodes and plant interactions

In Chapter 3, I investigated the effects that nematode infection can have on the growth and development of tomato, S. lycopersicum. There have been several crop loss studies conducted over the past several decades (some of which I have previously reported in Chapter 3); however, a standardised level of inoculum has not been determined or proposed. This issue is discussed further in the Future Work section of this chapter. In my study using 5000 nematodes per plant, I found that some growth parameters were affected even within a period of only one week after inoculation. While there were some parameters affected by both nematode species, other parameters were species-specific. For example, both M. javanica and P. thornei reduced shoot length and dry weight of shoots and roots, irrespective of the plant growth stage at which the nematodes were applied. However, only M. javanica reduced root dry weight, the diameter of the largest mature fruit and fresh and dry weight of fruit whereas only P. thornei reduced the number of green leaves, the number of flowers and buds at harvest and the moisture content of fruit.

Surprisingly, there were a number of parameters which were not affected by nematode infection; these were different for each nematode species. For example, M. javanica did not reduce the number of green leaves, number of flowers and flower buds at harvest, nor the fruit moisture content, and P. thornei did not affect dry weight of roots, diameter of the largest mature fruit, and the fresh and dry weights of fruit. These findings clearly point to differences in the nematode-plant interaction between the two nematode species. As the initial response to infection in Arabidopsis was similar for both nematode species (Chapter 2), and assuming that this is similar for tomato, I postulate that the differences in growth and development of nematode- infected tomato are a result of plant responses to the different ways the two nematodes damage the host, the subsequent nematode activity, the location of damaged host cells, and the steps taken by the host plant to repair this damage.

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It appears likely that following infection by nematodes the tomato plant tries to counteract the adverse effects of infection by first reducing the shoot length, so that the other parameters are not affected. Stunting of plants as a result of nematode infection has been a symptom reported commonly in field crops and other plants (Fielding, 1959; Nickle, 1991; Luc et al., 2005). Plant stunting can also be associated with infection by bacteria, viruses and the deficiency of trace elements (Agrios, 2005; Luc et al., 2005). Probably, one of the initial measures a plant adopts to cope with the stress due to infection is reducing shoot growth and, consequently, the dry shoot weight. This seems to be a general coping response not affected by the species of nematode infecting its root system.

Reduction in dry weight of roots is probably due to the slowing down of root growth in M. javanica-infected roots. Severe reduction in the rate of root growth was observed in Arabidopsis (reported in Chapter 2), where root hairs and xylem differentiated close to and just behind the root tip.

Similarly to reduction in shoot length, the reduction in fruit diameter may be another way the plant has of coping with stress. In tomato it is quite possible that if the fruit size is larger they are fewer in number, and if they are smaller they are greater in number. Benton Jones (2007) found that fruit size and number, and the total yield are related - the total yield is usually constant, while fruit size and number can change. Thus, the plant itself has the ability to regulate fruit size according to changing conditions. However, I did not check the number of seeds per fruit, which, ultimately, is probably the best indicator of a plant’s ability to propagate itself.

Moisture content of tomato fruit from plants inoculated with M. javanica was similar to fruit from uninfected plants. This indicates that water absorption and translocation within the plant was not significantly affected by the nematode damage. This conclusion appears to be supported by my observations in Arabidopsis (Chapter 2) that rapid repair of Meloidogyne-disrupted xylem tissue occurred. This suggests that the plant is still able to effectively absorb and translocate water and nutrients from the soil. This might also explain the corresponding lack of response in some other growth parameters such as the number of green leaves and flowers and buds.

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Compared to infection by M. javanica, P. thornei affected a greater number of growth parameters in plants. In tomato and Arabidopsis it was shown (Chapters 2 and 5) that the activity of P. thornei (migration and feeding) resulted in extensive tissue destruction within the root. The resulting stress from reduced absorption of water and nutrients from this damage may have resulted in the plant diverting its resources to the point of infection to combat the pathogen. However, in tomato, the plant seems to have compensated for direct tissue damage as there was no significant reduction in root dry weight. The fresh and dry weights of fruits were also not affected by nematode infection at the inoculum levels used in this experiment.

The reduction in percentage moisture content of fruit from Pratylenchus-infected plants suggests disruption in the uptake and conduction of water from roots to other parts of the plant. This is supported by the extensive tissue disruption observed in Pratylenchus-infected roots in Chapter 5. The disruption in water translocation occurred even though the nematode damage is mostly in the epidermis and the cortex, and the vascular bundle is left intact. This shows the importance of epidermal and cortical tissue in water uptake and translocation. In the case of infection by M. javanica, the vascular region is affected, and even then the water translocation is not affected significantly.

When I compared the percentage reduction of key growth parameters to the control, I found that inoculation with M. javanica reduced the dry weight of shoots and roots more than P. thornei. It was interesting to note that inoculation with M. javanica produced more flowers and buds than control plants while P. thornei-inoculated plants had significantly fewer flowers and buds at harvest. This may mean that either M. javanica induced early flowering or that P. thornei delayed the flowering in tomato.

Above results show that both M. javanica and P. thornei caused significant damage to tomato plants during the initial vegetative phase of the plant as well as the reproductive phase. Even a short period of one week of infection affected tomato growth. Therefore, any nematode control applied in the first few weeks of vegetative growth and later in the early reproductive phase will benefit the plant.

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6.1.3 Soil characteristics and plant growth parameters

In Chapter 4, I discussed my investigations on the movement of M. javanica and P. thornei towards the host root in three different soil texture types, and their subsequent infection of tomato roots. Both nematode species consistently increased in their numbers within roots up to ten days after inoculation in both the sand and clay soils. At the first assessment 3 DAI, the number of M. javanica in tomato roots was greater for those in clay than those in sand. In the sandy clay loam soil the highest number of nematodes in roots were observed at 3 DAI and declined up to 10 DAI, although they were consistently much lower than in the other two soils. I have suggested that this is probably due its chemical (and, possibly, biological) properties; having the lowest pH and the highest percentages of total organic carbon, total organic nitrogen, organic matter, and the lowest C : N ratio. It appears that these factors had a greater impact on nematode movement into roots than did its texture, for which it was originally chosen. Other workers have previously reported that organic matter improves plant growth in nematode-infected areas. It is probable that this factor (i.e. high organic matter content) was in play in my studies.

However, it should be noted that in the case of P. thornei, its transitory nature in roots underestimated its movement to roots in all cases, A number of tomato roots, were observed to have damaged cortical tissue, but no nematodes were observed within them.

Another interesting aspect is that sex determination in M javanica may be associated with environmental conditions (Triantaphyllou, 1971); this may possibly include soil characteristics. Thus, the soil textural and chemical characteristics, combined with other environmental conditions such as the relatively high experimental temperature, may have resulted in the formation of more males. Nematodes may have then left the roots, resulting in a low count. However, I did not investigate this.

Unlike M. javanica, the highest number of P. thornei at 3 DAI was found in roots of in tomato plants grown in sandy soil showing that for P. thornei this soil was most suitable for moving towards and infecting tomato roots. Sandy soil had the highest percentage component of sand; in the presence of sufficient moisture, the large pore

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size resulting from the larger particle size could have supported movement of this nematode, with its larger body diameter, towards the host root. Similar to observations in plants infected by M. javanica, the percentage counts of P. thornei in tomato roots grown in sandy clay loam soil were consistently low. The low population of P. thornei in plant roots observed in this experiment may be explained by the fact that sandy loam soil had, comparatively, the highest level of organic matter. P. thornei can survive in soil for a considerable time in the absence of host roots. Entrance holes and other tissue damage were frequently noted in inoculated roots, even in the absence of nematodes proving their prior entry into roots. The nematode-suppressing properties of the by-products of organic matter breakdown (such as ammonia), and the presence of antagonistic microorganisms, as has been previously described for M. javanica, may also explain the low nematode population in tomato roots grown in sandy clay loam soils.

Overall, results of this study indicated that many factors influence the movement of Meloidogyne javanica and Pratylenchus thornei towards tomato roots. Some of these factors include the soil characteristics such as texture, pore size and chemical composition, soil moisture and the diameter of the nematode body and its energy reserves.

In the experiment described in Chapter 3, an initial inoculum level of 5000 nematodes per plant was used, while in the experiment in Chapter 4, 300 nematodes per plant was used. The total number of nematodes ultimately infecting the root was not counted in Chapter 3. However, in Chapter 4, I found that only a very small portion of the initial inoculum was counted within roots (up to about 11% of the initial inoculum). Considering the fact that organic matter was a standard component in the potting mix used in Chapter 3, it may have an effect on the number of nematodes which actually reached the root. Thus, in the field if 5000 nematodes were present only a small percentage of these may ultimately reach and infect plant roots if organic matter was present. In addition, the soil texture and composition also would affect the number of nematodes actually able to move to and infect the plant.

In Chapter 4, I found that a large number of nematodes were found within the roots as early as three days after inoculation. The ability of the nematode to move and

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infect the plant within such a short time may be the reason for significant reduction of some growth parameters noticed even one week after inoculation in Chapter 3.

Castagnone-Sereno et al. (2006) proposed that a “fitness cost” may be associated with nematode virulence despite parthenogenetic reproduction. Highly virulent genotypes may develop within a generation, but their reproductive potential may be reduced. Such cost may impose a direct constraint on co-evolution between plant and the nematode populations, resulting in implications for the successful management of resistant cultivars in the field. Moreover, conditions prevalent in the field can be significantly different to the experimental conditions at any given time.

Many Australian soils are naturally low in nutrients and organic matter, and the soils used in this experiment were typical of these. However, there may be high variation in soil physical and chemical characteristics in farming situations. This variation mainly results from the initial soil formation as well as agricultural practices, such as tillage, fertiliser use and incorporation of organic matter. Based on my results, both physical and chemical factors may significantly affect nematode movement towards roots of potential host plants.

The results of this study support the view that incorporation of organic matter into soils may reduce the severity of plant parasitic nematode infection, even in soils that would otherwise be conducive to crop damage by nematodes. In combination with other integrated nematode management strategies, such as soil solarisation and/or crop rotation with non hosts, organic soil amendments could be considered to be an integral part. If successful, this could result in a reduction in environmental pollution by the reduction in the amount of toxic pesticide chemicals, increased crop yield, and, ultimately, increased farmers’ incomes.

6.1.4 Temporal study of nematode parasite – host plant relationships

In Chapter 5, I discussed a temporal study I undertook on the early infection process by M. javanica and P. thornei on tomato plants. As mentioned earlier, it was easy to distinguish the different life cycle stages in M. javanica; but this was difficult for P. thornei. I found significant similarities, as well as differences, between these two of

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nematode species with respect to the early stages of infection. Both nematodes were similar in that they mostly damaged single cells during entry, although there were a few cases where one nematode damaged more than one cell. Also, both nematode species appeared to prefer some roots over others for infection. The reason for this is unclear, but may be related to chemical cues produced by the plant and/or the nematodes associated with the initial infection.

On the other hand, they acted differently in several respects. There was minimal disruption in the surrounding host cells during migration of M. javanica, while there was extensive tissue disruption and discolouration in the case of migration by P. thornei. M. javanica was almost always observed migrating in between two cell walls, while P. thornei was observed mostly attacking the cross walls of cortical cells. The nematode may have been responding to signals from the end walls, which have different origin, structure and properties to the lateral walls (Collings and Wasteneys, 2005). Once P. thornei had entered the host I observed, in several cases, mostly at 6 DAI, that there were single nematodes coiled within a single cell. This was probably associated with the rest phases the nematode undergoes between feeding and migration, as previously observed by Zunke (1990) in root hairs.

Oval, highly stained, unidentified structures believed to be faecal matter were also observed, within roots infected by P. thornei, while no similar observation was made in roots inoculated with M. javanica. However, in M. javanica, nematodes moulted within the root and the shed cuticle was clearly observed in tomato roots, still attached to the nematode. Each moult was accompanied by a change in shape, where the nematode seemed to contract itself towards its head region. No such observation was made in P. thornei-infected tomato roots.

6.2 Usefulness / implication of my work to others

The techniques I utilised for the cytoskeletal study will be useful for further studies. Whole root immunolabelling can be used to study the initial plant responses to nematode infection in more detail. At the early, as well as later, stages when the galls become too thick for whole root studies, the BMM sectioning technique can be used to study these structures and their formation within giant cells and galls. Even though

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I used A. thaliana as the host for cytoskeletal studies, it helped to understand how other, natural host plants, such as tomato, are affected. Advances in nanoscience technology means that there is increased scope to study as yet unexplained aspects of early host-pathogen interactions. Such studies are now possible, as the basic methods and protocols have been developed and established during the course of my project. If better culture techniques to obtain aseptic P. thornei can be developed, the present techniques could be used to study its effects on the cytoskeleton in more detail, using a more suitable host such as tomato. For this, BMM resin embedding and sectioning followed by immunolabelling will be a more suitable method, especially in the late stages of gall development, because of limitations of the microscope to handle thick tissue samples such as nematode galls.

My pot culture experiments demonstrated that infection by nematodes, even for only a week, can adversely affect plant growth. My investigations used single nematode species. Under field situations, there may be several species of plant nematodes present as well as other plant pathogens. Therefore, the damage to plants would probably be even more that what I observed in my investigations. As discussed above, application of organic matter, spread over different stages for the duration of the crop, may improve growth and yield, by reducing the impact of nematodes. The beneficial effects of increased levels of organic carbon and nitrogen have been emphasised by the results of my investigations.

A better understanding of the mode of infection and the plant response may, in future, help to develop more resistant varieties or root stocks for improving nematode management in different soils. As soil texture and composition also significantly affect infection, combined or integrated pest management strategies could be developed to maximise yield, not only in tomato, but also in other crops.

6.3 Integrated management of nematode pests

A number of nematode control methods are presently available: these are discussed in more detail in Chapter 1. The results of my study suggest that nematode control strategies could be developed for field crops based on manipulation of soil structure, chemistry and organic matter content; and these strategies could be spread over the

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duration of the crop. However, for maximum benefit, they would be best if targeted at the vegetative and early reproductive stages of the plants. Barker and Koenning (1998) have previously suggested that an integrated approach, involving combinations of resistant cultivars, crop rotation, biocontrol agents, and clean and selected cultural practices together with the judicious use of nematicides, is likely to be the most effective in controlling plant parasitic nematodes. This is likely to be the case for management of M. javanica and P. thornei. The main objective of this approach is to keep the populations of plant parasitic nematodes below economic threshold levels (Bird, 1987; Sikora et al., 2005), and not their complete eradication.

6.4 Impacts of climate change on plant parasitic nematodes

Global warming has serious implications for farming worldwide. Widespread droughts and increases in atmospheric temperature can create conditions either favourable or detrimental to plant parasitic nematodes such as M. javanica and P. thornei. General predictions include high frequency of extreme high temperatures, increased precipitation in summer and reduced rainfall during winter, resulting in conditions favourable for plant pathogens world wide, including Australia (Garrett et al., 2006). The predictions for climate change in Australia suggest that annual average temperatures over most of the country will rise by 0.4-2.0° C by 2030, and that annual average rainfall in the north should remain similar to current levels or even increase, whereas the south-east region will decrease (range -10 to + 5%) over the same period (Whetton, 2001).

Apart from their direct effect on pathogens, these predicted changes may also cause water stress in dryland crops, and/or limited water for irrigation, which may result in breaking of their resistance to pathogens, and increasing their susceptibility to pests and diseases. These are likely to have both direct and indirect effects on nematode- host interactions. M. javanica and P. thornei infect crops grown in preferably warm, partly dry, but moist, soils with a relatively high sand content and low organic matter. However, any negative impacts of climate change could be counteracted by using modelling to develop adaptive changes in land use patterns. The results from my study may provide some, albeit limited, assistance to farmers in dealing with the effects of climate change. However, more focussed studies would

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have to be carried out to elucidate the complex interactions between nematodes and their plant hosts under scenarios of climate change. Given the importance of root parasitic nematodes and wide range of soil types in world crop production systems, this study indicates that these relationships need to be explored further in both laboratory and field investigations.

6.5 Future research work

• The findings from Chapter 2 require further investigation. Now that I have shown that these techniques are useful, more frequent observations than I conducted will elucidate the cytoskeletal changes beyond what I have achieved.

• The cytoskeletal study was undertaken in Arabidopsis as the host plant. The results of my (initial) studies indicate that the same results are likely to be observed in tomato. However, further studies should be conducted with tomato and Arabidopsis plants, to enable a more direct comparison.

• Further investigations with my two nematode species are required to determine the inoculum level required to reach the damage threshold in pot trials. This has been determined for a number of crops grown in field soils in some locations; however in pot trials, workers have used several different levels in a range of studies.

• I have postulated several reasons why in soils with high organic matter the lowest number of nematodes reached plant roots. However, further investigations need to be undertaken to identify the factor or combination of factors responsible for this.

• Further studies should be conducted to determine the extent of the relationship between the populations of Pratylenchus thornei in roots vs. their level in the soils in which plants are growing, in the presence of both high and low organic matter. This is important because, although I observed that

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roots were damaged in Pratylenchus-inoculated tomato plants, I did not record any nematodes within the roots in sandy clay loam soil, which had the highest organic matter content.

• Despite the work reported in this thesis, the relationships between the damage caused by the two species of nematodes and their effects on the physiology and subsequent growth and development of the host plants require will require extensive further study.

6.6 Final conclusion

Plant parasitic nematodes cause important changes in the hosts they infect. These changes start at the cellular level. The initial interaction between the parasite and the host may be similar; however, in later stages there are significant differences in the host response to different nematode species, which can ultimately affect plant growth and development.

Nematode damage can be minimised in the field by manipulating the growing environment, such as soil characteristics and the timing of application of organic amendments. It is necessary to consider the role climate change will play in future of the worldwide agricultural industries, especially in Australia.

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APPENDIX 1 RESULTS OF ANALYSIS OF SOILS USED IN POT TRIAL

Texture - Heavy Clay Average Stdev Saturation percentage SP% 61.8 1.3 Clay % 60.6 2.0 Sand % 33.4 0.2 Silt % 6.0 2.0 Electrical conductivity EC 1:5 dS/m 0.2 0.0 Electrical conductivity ECe uS/cm 1306.0 34.0 Electrical conductivity (saturated paste) ECe dS/m EC1:5 Conversion factor to Ece 5.5 0.2 pH Water 6.5 0.3 pH saturated extract 6.8 0.3 Spectrophotometer UV 254 nm 0.7 0.1 254nm Filtered samples as dissolved organic carbon 0.2 0.0 Relative particulate turbidity > 0.45uM 0.4 0.1 254nm extract 0.5 0.0 Total organic carbon TOC % 0.9 Organic matter OM % 1.5 Total organic nitrogen TON % 0.05 C:N ratio 17.6:1

Texture - Sand Average Stdev Saturation percentage SP% 19.2 1.4 Clay % 8.0 1.2 Sand % 91.3 0.0 Silt % 0.7 1.2 Electrical conductivity EC 1:5 dS/m 0.1 0.0 Electrical conductivity ECe uS/cm 1928.3 7.5 Electrical conductivity (saturated paste) ECe dS/m 1.9 0.0 EC1:5 Conversion factor to Ece 14.3 0.4 pH Water 6.1 0.2 pH saturated extract 5.0 0.1 Spectrophotometer UV 254 nm 2.2 0.1 254nm Filtered samples as dissolved organic carbon 0.3 0.1 Relative particulate turbidity > 0.45uM 1.9 0.1 254nm extract 0.7 0.0 Sample Prep Total organic carbon TOC % 0.6 Organic matter OM % 1.0 Total organic nitrogen TON % 0.0 C:N ratio 18:1

Texture - Sandy Clay Loam Average Stdev Saturation percentage SP% 49.6 1.2 Clay % 22.6 0.2 Sand % 61.5 0.2

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Silt % 15.9 0.2 Slaking 1.0 0.0 Electrical conductivity EC 1:5 dS/m 283.0 1.2 Electrical conductivity ECe uS/cm 2.9 0.3 Electrical conductivity (saturated paste) ECe dS/m EC1:5 Conversion factor to Ece 10.1 1.0 pH Water 5.0 0.2 pH saturated extract 4.9 0.2 Spectrophotometer UV 254 nm 2.1 0.2 254nm Filtered samples as dissolved organic carbon 0.2 0.0 Relative particulate turbidity > 0.45uM 1.9 0.2 254nm extract 1.9 0.1 Sample Prep Total organic carbon TOC % 3.5 Organic matter OM % 0.6 Total organic nitrogen TON % 0.3 C:N ratio 11:1

Sample ID TOC% OM% TON% C:N ratio Sand (S) 0.6 1.0 0.03 18 is to 1 Sandy Clay Loam (SCL) 3.5 6.0 0.31 11 is to 1 Heavy Clay (HC) 0.9 1.5 0.05 17.6 is to 1

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