Biology and Molecular Characterisation of the Root Lesion Nematode, Pratylenchus Curvicauda

Biology and Molecular Characterisation of the Root Lesion Nematode, Pratylenchus curvicauda

This thesis is presented by

FARHANA BEGUM

For the degree of Doctor of Philosophy

School of Veterinary and Life Sciences,

WA State Agricultural Biotechnology Centre (SABC),

Murdoch University, Perth, Western Australia

July 2017

Declaration

I declare that this is my own account of my research and contains as its main content, work which has not previously been submitted for a degree at any tertiary educational institution.

FARHANA BEGUM

Abstract

Australia is the driest inhabited continent with about 70% of the land arid or semi- arid, and soils which are geologically old, weathered, and many are infertile. This is a challenging environment for agricultural production, which is further impacted by biotic constraints such as root lesion nematodes (RLNs), Pratylenchus spp. These soil-borne nematodes cause significant economic losses in yields of winter cereals, and in other crops, particularly under conditions of moisture and nutrient stress. RLNs are widely distributed in

Australian broadacre cropping soils, and losses in cereal production are greater when more than one RLN species is present, a situation which often occurs in Western Australia (WA).

Hence, to develop appropriate management regimes, accurate identification of RLN species is needed, combined with understanding the biology of host-nematode interactions.

The initial aim of this research was to extend the molecular and biological characterisation of P. quasitereoides, a recently described species of root lesion nematode from WA. Morphological measurements of two important characters, tail shape and the per cent distance of the vulva from the anterior end of the nematode body, were made from nematodes collected from the four locations of WA. These measurements did not match with the published data on P. quasitereoides. As a result, nematode samples from Pingelly were sent to Dr. M. R. Siddiqi in the UK for detailed morphometric measurements: he concluded that the nematodes from Pingelly differed from other recognised species of Pratylenchus in

Australia. The longer conus (more than half of the spear length), three annules in the lip region, large rounded basal knobs, phasmids in the anterior region of the tail, usually about one-fourth the tail length behind the anus, and the absence of males were good characters for species comparisons. The morphometric measurements suggested that the species under study was in fact P. curvicauda, a species which had been reported previously and described

iii from WA (Siddiqi et al., 1991). Analyses of SEM images also supported the morphological description of the nematode samples from Pingelly as being P. curvicauda.

Study of the genetic variation of this species with other known species of

Pratylenchus was undertaken based on analysis of sequences of both the ribosomal DNA

(rDNA) ITS and D2-D3 regions. Sequence analyses showed that there was considerable variation in P. curvicauda rDNA in samples from four wheatbelt regions, and this was also the case for P. neglectus, P. penetrans and P. thornei. The ITS sequences of P. curvicauda differed by 39% from those of P. neglectus and P. thornei, but were more similar to those of

P. penetrans. Nucleotide variations between sequences of P. curvicauda and those of P. neglectus, P. penetrans and P. thornei were mainly in the ITS1 and ITS2 regions, which differed by 44% and 49% in nucleotide sequences of these regions with P. neglectus and P. thornei, whereas with only 10% in both regions compared to P. penetrans sequences. For the

D2-D3 region, sequences from P. curvicauda, P. neglectus, P. penetrans and P. thornei differed by up to 30% at the nucleotide level. The D2-D3 sequences of P. curvicauda were very different from the published sequences for P. quasitereoides, with up to a 39% variation in nucleotide sequence. These results support the morphological data that Pratylenchus sp. collected from the four WA locations did not appear to be any of the species of Pratylenchus currently described as being in WA (i.e. P. neglectus, P. penetrans, P. thornei or P. quasitereoides). This result prompted a more detailed study of the biology of the nematode.

The ability of different plant taxa/genotypes to support the reproduction of P. curvicauda was assessed using soil collected from Pingelly under glasshouse conditions.

Crop and pasture species challenged included wheat, barley, pasture species, brassicas, and model plants, both by manual inoculation and grown in naturally infested field soil. Wheat, barley and sorghum were good hosts; canola, tomato and other brassicas were poor hosts; most pastures species and tobacco were non-hosts.

A part of this research focused on how P. curvicauda was attracted to and penetrated roots of host plants identified as resistant or susceptible. Since attraction to and entry into roots of hosts classified as resistant or susceptible was essentially the same, host resistance is probably expressed later in the interaction. The life cycle of this species was also studied. It took 35-42 d to complete one generation in susceptible wheat roots under glasshouse conditions.

Histological studies of roots infested by P. curvicauda were also undertaken. Cellular changes in infested roots included deformation of cell wall, loss of cytoplasmic contents and death of invaded cells, and there was some evidence of changes in cells further from the invading nematodes. There were no clear differences in the damage caused in susceptible and resistance hosts, at least up to 7 d after infection.

This study is the first to investigate the biology and molecular characteristics of P. curvicauda.

Table of Contents

Declaration...... ii Abstract ...... iii Table of Contents ...... vi Abbreviations ...... x List of Figures and Tables ...... xii Publications and Presentations ...... xv Acknowledgements ...... xvii Chapter 1. Introduction and literature review...... 1 1.1 General introduction ...... 1 1.2 Distribution and economic importance of Root Lesion Nematodes (RLNs) ...... 2 1.3 Morphology of RLNs ...... 5 1.4 Molecular diagnostics and genetic diversity in RLNs ...... 10 1.5 Culturing, extraction and quantification of RLNs ...... 11 1.6 Edaphic factors which affect RLNs ...... 13 1.7 Biology of RLNs ...... 14 1.8 Host range and pathogenicity of RLNs ...... 16 1.9 Host location and feeding of RLNs ...... 17 1.10 Histopathology and ultra structural changes in host cells infested with RLNs ...... 18 1.11 Molecular studies of PPN-plant interactions ...... 18 1.12 Conclusion ...... 20 1.13 Aims and Objectives ...... 20 Chapter 2. Morphological characterisation of an undescribed Pratylenchus species from Western Australia ...... 22 2.1 Introduction ...... 22 2.2 Materials and methodology ...... 23 2.2.1 Extraction and culturing of nematodes ...... 23 2.2.2 Fixing nematode samples for measurement ...... 24 2.2.3 Preparation of nematode samples for Light Microscopy...... 24 2.2.4 Preparation of nematode samples for Scanning Electron Microscopy (SEM) ...... 24 2.3 Results ...... 25 2.3.1 Morphological features and measurements of nematode samples ...... 25 2.3.2 Detailed description and morphometric analyses of nematode samples from ...... 28 Pingelly ...... 28 vi

2.3.3 Comparative morphological features of the undescribed species from Pingelly and its relationship with other species of Pratylenchus ...... 30 2.3.4 Features of nematode samples using Light and Scanning Electron microscopy ..... 33 2.4 Discussion ...... 37 2.5 Conclusion ...... 38 Chapter 3. A study of genetic variation in Pratylenchus spp. in cereal growing regions of Western Australia ...... 39 3.1 Introduction ...... 39 3.2 Materials and methods ...... 40 3.2.1 Nematode populations, extraction and culture ...... 40 3.2.2 DNA extraction ...... 42 3.2.3 PCR amplification, cloning and sequencing...... 43 3.2.4 Analyses of sequences ...... 44 3.3 Results ...... 45 3.3.1 Morphology of the Pratylenchus species studied ...... 45 3.3.2 Length of ITS sequences of P. curvicauda, P. neglectus, P. penetrans and P. thornei ...... 45 3.3.3 Variation in ITS sequences of P. curvicauda ...... 46 3.3.4 Variation in ITS sequences between P. curvicauda, P. neglectus, P. penetrans and P. thornei ...... 46 3.3.5 Comparison of ITS sequences of RLNs from WA and global data ...... 47 3.3.6 Analyses of D2-D3 sequences ...... 48 3.3.7 Comparison of D2-D3 sequences of P. curvicauda with those of P. quasitereoides ...... 49 3.3.8 Comparison of D2-D3 sequences ...... 50 3.3.9 Phylogenetic analyses of ITS and D2-D3 sequences ...... 51 3.4 Discussion ...... 59 3.5 Conclusion ...... 61 Chapter 4. Susceptibility of cereals, common crops and pasture species to P. curvicauda ...... 62 4.1 Introduction ...... 62 4.2 Materials and methods ...... 63 4.2.1 Sources of plants and germplasm ...... 63 4.2.2 Growing test plants and nematode inoculation...... 64 4.2.3. Observation of inoculated plants and nematode infestation analysis ...... 66 4.3 Results ...... 67

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4.3.1 Reproduction of P. curvicauda in test plants inoculated with vermiform stages of nematodes ...... 67 4.3.2 Challenge of wild germplasm of cereals with soil naturally infested with P. curvicauda ...... 71 4.3.3 Plant response, root structure and nematode development ...... 75 4.4 Discussion ...... 75 4.5 Conclusion ...... 78 Chapter 5. Attraction, penetration and biology of P. curvicauda ...... 79 5.1 Introduction ...... 79 5.2 Materials and methods ...... 80 5.2.1 Nematode inoculum ...... 80 5.2.2 Attraction of P curvicauda to host ...... 81 5.2.3 Penetration of P curvicauda to host ...... 81 5.2.4 Life cycle of P curvicauda ...... 82 5.2.5 Staining and counting of nematodes ...... 83 5.3 Results ...... 83 5.3.2 Attraction of nematodes to resistant and susceptible roots...... 83 5.3.1 Penetration of nematodes in resistant and susceptible genotypes ...... 85 5.3.3 Life cycle of P. curvicauda in wheat ...... 87 5.4 Discussion ...... 90 5.5 Conclusion ...... 93 Chapter 6. Histological responses of susceptible and resistant hosts infested with P. curvicauda ...... 94 6.1 Introduction ...... 94 6.2 Materials and methods ...... 95 6.2.1 Preparation of nematode inoculum ...... 95 6.2.2 Growing of plants and inoculation of nematodes ...... 95 6.2.3 Staining of nematodes ...... 95 6.2.4 Harvesting and fixation of roots for light microscopy (LM) and transmission electron microscopy (TEM)...... 96 6.3 Results ...... 97 6.4 Discussion ...... 104 6.5 Conclusion ...... 107 Chapter 7. General discussion ...... 108 7.1 Morphological features of the undescribed Pratylenchus species from four paddock soils of WA...... 109

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7.2 Genetic variation of P. curvicauda with other Pratylenchus populations in WA ...... 109 7.3 Host preferences of P. curvicauda ...... 111 7.4 Biology of P. curvicauda ...... 112 7.5 Future recommendations ...... 116 Appendix ...... 118 References ...... 121

Abbreviations

AR Arthur River bp base pair CMCA Centre for Microscopy, Characterisation and Analysis CN cyst nematode cv. Cultivar DAFWA Department of Food and Agriculture, Western Australia dai days after inoculation d days DNA Deoxyribo Nucleic Acid FDA Fluroscein Diacetate FITC Fluorescein Isothiocyanate HIGS Host-Induced Gene Silencing h hours IEF Iso-Electric Focusing ITS-RFLP Internal Transcribed Spacer- Restricted Fragment Length Polymorphism J1 Juvenile stage 1 J2 Juvenile stage 2 J3 Juvenile stage 3 J4 Juvenile stage 4 KAT Katanning LCM Laser Capture Microdissection LM Light Microscopy MAD-PCR Multiplex Anti-primer Denaturation-Polymerase Chain Reaction MALDI-TOF Matrix Assisted Laser Desorption Ionisation-Time Of Flight ml millilitre min minute NCBI National Centre for Biotechnology Information NSW New South Wales nt nucleotide

PCR Polymerase Chain Reaction PN Pingelly PPN Plant Parasitic Nematode QPCR Quantitative Polymerase Chain Reaction QTL Quantitative Trait Locus RAPD Random Amplified Polymorphic DNA rDNA ribosomal DNA Rf Reproduction factor RFLP Restricted Fragment Length Polymorphism RKN Root-knot nematode RLN Root Lesion Nematode RNAi Ribonucleic acid-interference SCAR-PCR Polymerase Chain Reaction SDS-PAGE Sodium Dodecyl Sulphate Polyacrylamide Gel Electrophoresis SEM Scanning Electron Microscopy s second TEM Transmission Electron Microscopy WA Western Australia µl Microliter µm Micrometer

List of Figures and Tables

Figures Page No. Figure 1.1. Geographical distribution of RLNs worldwide (Castillo & Vovlas 2007). 3

5 Figure 1.2. Distribution of RLN species in Australia. (Source: http://www.pir.sa.gov.au/research/services/molecular_diagnostics/predicta_b)

7 Figure 1.3. Adult female of P. curvicauda (see Chapter 2) (left) and a male of P. penetrans (right).

Figure 1.4. Life cycle of RLNs (Jones & Fosu-Nyarko 2014). 15

Figure 2.1. Morphology of the undescribed root lesion nematode from Pingelly; 26 a. Adult female, b. head region, c. tail region (scale bar=100 μm).

30 Figure 2.2. Line drawings of different body parts of the undescribed Pratylenchus sample from Pingelly, WA: a-head, b-oesophageal region, c- vulva region, d & e-tail ends (as illustrated by Dr. M. R. Siddiqi, unpublished).

Figure 2.3 Differences in tail shape and relative position of the vulva of the undescribed sample 33 labelled as Pratylenchus sp. and three other known species (P. penetrans, P. neglectus and P. thornei) examined using a compound microscope (moving nematodes).

Figure 2.4. Light micrographs of different species of Pratylenchus from WA. a. Entire female 34 body (left), b. head (up) and c. tail region (bottom) of undescribed Pratylenchus from Pingelly; d. entire female body, e. head (up) and f. tail region (bottom) of P. neglectus; g. entire female body, h. head (up) and i. tail region (bottom) of P. penetrans; j. entire female body, k. head (up) and l. tail region (bottom) of P. thornei. (scale bar: a-c=50 µm and d-l=100 µm).

Figure 2.5. Female of undescribed Pratylenchus sp. from Pingelly under SEM. a. a whole 35 nematode body, b-c. enface view, d. lateral field at vulval region.

Figure 2.6. Female of undescribed Pratylenchus sp. from Pingelly under SEM. a. tail region, b. 36 lateral field at middle of the body, c. tail terminus (arrow points to anus), d. vulval region (arrow points to vulva).

Figure 3.1 The four locations from where Pratylenchus samples were collected. 41

43 Figure 3.2. The location of the primers used to amplify different regions of rDNA genes of Pratylenchus spp.

Figure 3.3. Amplified ITS region of Pratylenchus spp.; 1. P. curvicauda, 2. P. neglectus, 3. P. 45 penetrans, 4. P. thornei, 5. Negative control; M=100 bp marker.

Figure 3.4. Amplified D2-D3 gene segments of Pratylenchus spp.; 1. P. curvicauda, 2. P. 48 neglectus, 3. P. penetrans, 4. P. thornei, 5. Negative control, M=100 bp marker.

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Figure 3.5. Multiple alignments of sections of D2-D3 with variation in 17 sequences of P. 50 curvicauda and published D2-D3 sequences of P. quasitereoides: AR- P. curvicauda sequences from Arthur River, WL- P. curvicauda sequences from Williams, PN- P. curvicauda sequences from Pingelly, KAT- P. curvicauda sequences from Katanning and D2-D3 sequences of P.quasitereoides from Hodda et al. (2014), as marked with the accession number from KM248297.1-248306.1. The numbers denote the number or identity of clones from the same location).

Figure 3.6. Phylogenetic relationships between ITS sequences of P. curvicauda, P. neglectus 52 (Pneg), P. penetrans (Ppen) and P. thornei (Pth), as inferred from Maximum Likelihood based on GTR+G+I model. Posterior probabilities from 50% are given for appropriate clades. AR- P. curvicauda sequences from Arthur River, WL- P. curvicauda sequences from Williams, PN- P. curvicauda sequences from Pingelly, KAT- P. curvicauda sequences from Katanning. Numbers are identifiers for clones.

54 Figure 3.7. Maximum Likelihood (GTR+G) tree using the ITS sequences of Pratylenchus spp. Phylogenetic analyses were conducted in MEGA6. The P. curvicauda sequences from the four locations of WA obtained in this study are labelled with a black dot. The accession number and details of the ITS sequences other Pratylenchus spp are listed in Appendix 1.

55 Figure 3.8. a. Maximum Likelihood (K2+G+I) tree constructed from the ITS of P. neglectus WA population (Pneg1.1-Pneg3.3) with corresponding sequences from GeneBank. Evolutionary analyses were conducted in MEGA6; b. Molecular Phylogenetic analysis by Maximum Likelihood method (T92+G) showing relationships among global population of Pratylenchus penetrans. Ppen1.1-Ppen5.2 is P. penetrans form WA.

Figure 3.9. Molecular phylogenetic analysis of ITS sequences of P. thornei by Maximum 56 Likelihood method (K2: Kimura 2-parameter), Pth1.1-Pth2.2 indicates P. thornei from WA.

Figure 3.10. Phylogenetic relationships of P. curvicauda sequences from the four locations in 57 WA and sequences of P. quasitereoides from Hodda et al., (2014) as inferred from D2-D3 sequence alignment under K2+G model. P. quasitereoides from Hodda et al., (2014), as marked with the accession number from KM248297.1-248306.1, named as Pratylenchus sp. Posterior probabilities more than 50% are given for appropriate clades. Newly obtained sequences of P. curvicauda from this study are indicated by a black dot.

Figure 3.11. Comparison of D2-D3 sequences from Pratylenchus spp. as inferred from 58 Maximum Likelihood using K2+G model. The P. curvicauda sequences from the four locations of WA obtained in this study are labelled with a black dot. P. quasitereoides from Hodda et al. (2014), as marked with the accession number from KM248297.1-248306.1) named as Pratylenchus sp.

69 Figure 4.1. The average number of P. curvicauda in soil and roots of crop and pasture plants 10 weeks after inoculation with a suspension of 1000 mixed stages of the nematode. Replicates (n=12).

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70 Figure 4.2. Multiplication of P. curvicauda in crops and pasture species in soil inoculated with standard nematode inoculum. 73 Figure 4.3. Population of P. curvicauda in roots of cereal germplasms in naturally infested soil (n=12).

74 Figure 4.4. Multiplication of P. curvicauda in crops and pasture species in naturally infested soil. Figure 5.1. Attraction of P. curvicauda to host roots in a Petri dish (R=Root, N=Nematode). 82

Figure 5.2. Seedlings in corrugated trays (left and middle) and transferred seedlings in tray pot 83 (right).

Figure 5.3. Attraction of nematodes to roots: a. movement of nematodes towards root hair zone 84 and upper part of roots after 6 h; b. nematodes around a root after 6 h (wheat cv. Calingiri); c. nematodes around a root after 2 h (AGG17458WHEA1).

Figure 5.4. Nematodes attracted towards roots of cereals and pastures after 2, 4, 6 and 12 h. The 85 designations are: S-susceptible, R-resistant, HS-highly susceptible, HR- highly resistant (refer to Chapter 4). For each time period, bars with the same letter did not differ significantly (P=0.05) according the Student's t-test.

Figure 5.5. Nematodes inside roots of wheat (AGG17458WHEA1) (left) and triticale 86 (AGG3959) (right) after 14 dai, staining with acid fuchsin.

Figure 5.6. Percentages of P. curvicauda in roots of cereals and pastures species 48 h to 14 dai. 87 The designations are: S-susceptible, R-resistant, HS-highly susceptible, HR- highly resistant (refer to Chapter 4). For each time period, bars with the same letter did not differ significantly (P=0.05) according to the Student's t-test.

Figure 5.7. Days required for J2-J3 of P. curvicauda to complete its life cycle in a susceptible 88 wheat cv. Calingiri (not to scale).

Figure 5.8. The average body length of the nematode P. curvicauda in wheat cv. Calingiri roots 88 at different days after inoculation.

Figure 5.9. Life stages of P. curvicauda (J2-J2) in wheat roots stained with acid fuchsin from 15- 89 50 dai.

Figure 5.10. Brown discoloured wheat roots at 55 dai; b-d: Fungal structures found in roots of 90 wheat cv. Calingiri.

Figure 6.1. P. curvicauda in wheat cv. Calingiri roots at different times after inoculation: a. 3 dai; 97 b. 7 dai; c. 15 dai.

Fig. 6.2. Nematodes lying on cortical cells in: a. Longitudinal paraffin section of wheat cv. 98 Calingiri roots at 3 dai; b. wheat cv. Calingiri roots at 7 dai; c. barley cv. Hamelin roots at 7 dai; d. wild barley roots at 7 dai. (N=nematode).

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Fig. 6.3. Light micrographs of nematodes in the cortex of wheat roots at 3 dai: a. P. curvicauda 98 in cortical parenchyma (p=pericycle, e=endodermis, nu=nucleus, N=nematode), deformed endodermis, thickened cell walls in surrounding cells (cross section); b. longitudinal section of roots showing collapse of cytoplasm (marked with arrow), and enlarged nuclei (longitudinal section).

Figure 6.4. a & b. Paraffin sections of wheat cv. Calingiri roots at 3 dai showing intra-cellular 99 migration of P. curvicauda (nu=nucleus; N=nematode); c. control roots without nematode (longitudinal sections).

Fig.6.5. Nematodes in glutaraldehyde/OsO4 fixed roots under a dissecting microscope. a. wheat 100 cv. Calingiri 3 dai; b. wheat cv. Calingiri at 7 dai; c. Barley cv. Hamelin at 7 dai.

Figure 6.6. Light and electron micrographs of infested wheat (cv. Calingiri) roots at 3 dai. a. LM 101 of infested roots; b. TEM showing cells containing nematodes (N), but otherwise cells devoid of cytoplasmic contents; c. deformed and compressed phenotype of adjacent cells; d. granular material in the cytoplasm, larger cavities (V) and tannin (t) deposition in infested root cells.

Figure 6.7. TEM of wheat (cv. Calingiri) roots at 7 dai infested with P. curvicauda a & b. 102 adjacent cell devoid of cytoplasmic content, deformed plasma membrane (p= pericycle, e= endodermis, N= nematode); c. deformed endodermis, nucleus with condensed chromatin; d. cross section of a control root without nematode infection.

Figure 6.8. Barley (cv. Hamelin) roots at 7 dai infested with P. curvicauda. a. light micrograph 103 of root; b. nematode inside cell showing devoid of cytoplasmic content with disorganised cytoplasm in adjacent cell; c. ruptured epidermal cell wall (as indicated by arrow) (N=nematode).

Figure 6.9. Wild barley roots at 7 dai infested with P. curvicauda. a & b. compressed cells with 104 degeneration of cell contents; c. nematode showing vulva; d. control cells without nematode infestation.

Figure 7.1. Interaction of RLNs with host cells. Activities of effectors, proteolytic enzymes, and 115 other proteins during migration and feeding by nematode in host cells (Fosu-Nyarko & Jones 2016).

Tables:

Table 1.1. Comparative morphometric measurements of some Pratylenchus spp. 9

Table 1.2. Examples of the effects of host species and temperature on completion of RLN life 15 cycles 27 Table 2.1. Comparative morphometric measurements of Pratylenchus spp. adult female from WA

32 Table 2.2 Comparative morphometric measurements of undescribed Pratylenchus from Pingelly with other closely related species

65 Table 4.1. Crops/cultivars assayed for supporting the reproduction of P. curvicauda

Publications and Presentations

Begum, F., Fosu-Nyarko, J., Sharma, S., Macleod, B. & Jones, M.G.K. (2016). ITS sequences reveal genetic variation within populations of ‘Pratylenchus quasitereoides’ and between Pratylenchus species in Australia. Proceedings of 32nd Symposium, European Society of Nematologists (ESN), Braga, Portugal, Aug 28- Sept 1, p275.

Begum, F., Sharma, S., Jones, M.G.K., Macleod, B. & Fosu-Nyarko, J. (2015). Host preferences of ‘Pratylenchus quasitereoides’ - a new challenge in soil-root battlefield. Australasian Plant Pathology Society (APPS) Conference, Fremantle, Perth, 14-26 September, p120.

Murdoch School of Veterinary and Life Sciences Poster Day 2016

Begum, F., Fosu-Nyarko, J., Sharma, S., Macleod, B. & Jones, M.G.K. (2016). ITS sequences reveal genetic variation in Pratylenchus species in Australia. Poster: 09 Nov 2016.

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Acknowledgements

First I would like to sincerely thank my principal supervisor Professor Michael G. K. Jones for encouragement and furthering my interest in Nematology. I would like to thank you for all of your support from the beginning of this long journey, for your patience, advice and for correcting this manuscript.

My sincere thanks go to Dr. Shashi Sharma and Bill Macleod, for giving me the opportunity to undertake a PhD with their initial support.

I am extremely grateful to Dr. John Fosu-Nyarko for useful advice given on many different aspects of my research. Thank you for encouraging and inspiring me to complete this PhD.

I thank Dr. Sarah Collins and the nematology group, DAFWA for giving me the support to start my nematode work, and for collecting nematode infested soil for my research. Thanks also to Helen Hunter for teaching me how to pick single nematode.

I would also like to thank all staff and researchers in the SABC. Thanks to Dr. Dave

Berryman for his support for lab equipment, to Frances Brigg for helping with DNA sequencing and its interpretation. I also thank Mr. Gordon Thomson for his assistance in histology, SEM and TEM sectioning, and to Mr. Michael Platten, DAFWA for helping me with TEM imaging.

Special thanks are reserved for all fellow students in the Plant Research Biotechnology group

(PBRG) working together in the lab, for having fun and for the wonderful time spent together.

To my family and friends for their love and continuous support. I acknowledge the support, understanding and patience of my beloved daughter AUDRI, without your cooperation I would not have been able to undertake this thesis. Love you Maa.

I would also like to acknowledge the financial support of Murdoch university which allowed me to do this PhD research.

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Chapter 1. Introduction and literature review

1.1 General introduction

Nematodes are arguably the most numerous metazoans in soil and aquatic sediments (Abebe et al., 2011). Around 25,000 species of nematodes have been described, of which at least

4,100 species are parasites of plants (Decraemer & Hunt 2006). Because of changes in cropping patterns, new species are being described, and others which were previously regarded as benign or non-damaging are now emerging as more serious pests (Nicol et al.,

2011). Plant parasitic nematodes (PPNs) possess a mouth stylet, a sharp hollow, needle-like tube, which is used to pierce plant cell walls, secrete enzymes for host parasitism and ingest nutrients from the host (Castagnone-Sereno 2002; Trudgill 1991; Zunke 1990). Depending on the genus, nematodes generally feed on below-ground plant parts including tubers, bulbs and roots, while some PPNs (e.g. Anguina spp., Aphelenchoides spp.) feed on above ground plant parts including seeds, flowers, buds, leaves and stems. PPNs cause crop yield losses estimated at 14.6% in developing countries and 8.8% in developed countries (Nicol et al.,

2011). Because many species have broad host ranges, and PPNs attack all crop plants, they cause annual crop losses estimated at $150 billion worldwide (Bird & Kaloshian 2003).

However, losses caused by PPNs are often underestimated because the symptoms visible on aerial parts of plants can be confused with symptoms resulting from nutritional or water stress, or virus infection.

Root lesion nematodes (RLNs), Pratylenchus species, are one of the most important groups of PPNs. They are common in all agricultural regions of the world and reduce crop yields by migrating in and feeding on root tissues (Castillo & Vovlas 2007). Because of wide host ranges and distribution, RLNs are considered to be the third most important group of

PPNs after root-knot (RKN) and cyst nematodes (CN) (Jones et al., 2013; Sasser & Freckman

1987). RLNs parasitise and cause significant yield losses in most agriculturally important 1

crops including cereals, legumes, banana, vegetables, sugarcane, coffee and fruit trees

(Castillo & Vovlas 2007). They are polyphagous, obligate, migratory root endoparasites.

Pratylenchus species are known as root lesion nematodes because cells of the root tissues they feed on die and often develop characteristic necrotic lesions (Castillo & Vovlas 2007;

Vanstone et al., 1998). The damaged tissues impair water and nutrient uptake by the roots, resulting in poor plant growth and yield (Orion et al., 1984; Farsi et al., 1993). The invasion and migration of RLNs through root tissues and direct feeding from cells also facilitates subsequent secondary infection by fungi and bacteria (Powell 1971). Unlike RKNs and CNs,

RLNs do not form a long term feeding site in infected roots, and all motile stages can enter and leave host roots: this is one reason why their presence is less obvious than RKNs and

CNs, and their numbers are more difficult to quantify. In general, because RKNs and CNs are easier to work with than RLNs, and there is scientific interest in how the feeding cells of

RKNs and CNs (giant cells or syncytia respectively) form and function, they have been the subject of more detailed research on molecular characterisation and detailed biology, and until recently such studies had been comparatively neglected for migratory endoparasites like

Pratylenchus.

1.2 Distribution and economic importance of Root Lesion Nematodes (RLNs)

The genus Pratylenchus has 80 described species (Castillo & Vovlas 2007; Subbotin et al.,

2008). They seem to be distributed worldwide (Fig 1.1) with some widespread species of this genus, while others are geographically restricted to particular regions (Ryss 2002; Siddiqi

2000). The greatest level of diversity of the genus occurs in Asia, with 40 species reported

(Castillo & Vovlas 2007). The geographic location, introductions and environmental coniditions determine which subset of nematodes are crop parasites in Australia (Hodda &

Nobbs 2008). In Australia, at least 11 economically important root lesion nematode species

(P. brachyurus, P. coffeae, P. crenatus, P. goodeyi, P. jordanensis, P. neglectus, P.

penetrans, P. thornei, P. vulnus, P. zeae and P. teres) are present (Hodda & Nobbs 2008).

All of these economically important species are considered to be major pests of field crops and tree species, distributed over temperate and tropical regions (Thompson et al., 2010). The only Pratylenchus species thought to be endemic to Australia was initially reported as P. teres (Hodda & Nobbs 2008). However, recently another root lesion nematode species was descibed in WA which was similar to P. teres but had sufficient differences to justify the description of a new species, P. quasitereoides (Hodda et al., 2014): study of this endemic species was to be the subject of this thesis.

Figure 1.1. Geographical distribution of RLNs worldwide (Castillo & Vovlas 2007)

After Antarctica, Australia is the driest continent on earth, and this presents a challenging environment for agricultural production. In rain limiting environments, Figure 2.1. Geographical distribution of RLNs worldwide (Castillo & Vovlas 2007) nematodes can be one of the major limiting factors in grain production: this is indeed the case in Australia (Thompson et al., 2008). About 85% of Australia’s wheat and barley is grown in Figure 3.1. Geographical distribution of RLNs worldwide (Castillo & Vovlas 2007) the southern and western wheat belt, which extends more than 1400 km through southern

New South Wales (NSW), Victoria and the southern part of South Australia, and to the south westFigure region 4.1. of Geographical WA (Vanstone distribution et al., 2008). of RLNs worldwide (Castillo & Vovlas 2007)

P. neglectus and P. thornei are reported as the two most widespread species infecting wheat in Australia (Fig 1.2), and they can cause losses in yield of more than AUD 73 million and AUD 50 million per annum respectively (Murray & Brennan 2009). These two species often occur as mixed populations in southern Australia (Nicol 1996). P. neglectus has been found in almost all cropping environments of southern NSW, Victoria, South Australia (SA) and in WA (Ophel-Keller et al., 2008; Vanstone et al., 2008; Riley & Kelly 2002) with yield losses varying from 10-30% (Vanstone et al., 1998, 2002a, 2008a; Taylor 2000). This species is also recorded as a devastating species of wheat in Israel, Oregon and Washington State in

North America, and in Mexico, with resulting yield loss depending on host and infected crop species (Smiley 2010).

Yield losses from P. thornei infestation in the northern grain growing area of

Australia have been estimated at more than 50% in susceptible wheat cultivars (Thompson et al., 2008, 2010). Wheat yield loss in NSW, Queensland, South Australia and Victoria from P. thornei damage can range locally from 40-85% (Doyle et al., 1987; Thompson et al., 1993;

Taylor & McKay 1993; Eastwood et al., 1994). The preferred crop host of P. thornei is wheat, but it also infests other grains and legumes in Mediterranean environments and countries such as India (Nicol et al., 2003).

P. penetrans is also an economically important and cosmopolitan species, often prevalent in irrigated sandy soils (Smiley 2010). It reduces yield and/or quality of potato, carrot, pea, bean and ornamentals in Europe (Talavera et al., 2001; Pudasaini et al., 2007), and parasitises wheat and barley in Australia and Canada (Nicol & Rivoal 2007).

P. zeae mainly infects corn and tobacco, but it is also pathogenic on sorghum and soybean (Fortuner 1976), and is the major nematode pest of sugarcane in Queensland,

Australia (Blair et al., 1999a, 1999b). The species recently described as P. quasitereoides was reported to be a pest unique to WA (Hodda et al., 2014), occurring with P. neglectus in WA

crops. The RLN species present in WA soils (Fig 1.2) cause yield losses estimated at 15-50%

(Collins et al., 2013). In 2016, 63% of National Variety Trial (NVT) paddocks sampled in

WA were infested with RLNs, of which 83% of infested paddocks contained P. neglectus and

23% P. quasitereoides (Collins et al., 2017). P. neglectus was the most common RLN species followed by P. quasitereoides and P. penetrans.

Figure 1.2. Distribution of RLN species in Australia. (Source: http://www.pir.sa.gov.au/research/services/molecular_diagnostics/predicta_b)

1.3 Morphology of RLNs

The taxonomic designation of RLNs is based on morphological features of individual adult nematodes. The overall body shape of nematodes is determined by the pressure of its internal body fluids pushing against its strong, but flexible, outer cuticle. The cuticle comprises of a series of fine rings (annulations) which allow the cuticle to bend at any point along its body.

Depending on the species, adult Pratylenchus nematodes are 300-900 µm long and relatively stout (body length ÷ body width, usually 20 to 30). The head of the nematode can be recognised by the presence of a short, dark spear with basal knobs (the stylet) inside the tip of the head. The stylet is hollow (like a hypodermic needle) and can be extended from the head when used for penetrating plant cell walls and feeding from cell contents. The very outer tip of the nematode head above the stylet (called the lip region) is characteristically flat and darker in the genus Pratylenchus. In a relatively clear area just below the stylet can be seen the round, muscular pump or metacorpus, the metacorpus pumps substances up and down the oesophagus of the nematode. Just below the metacorpus is another relatively clear area that contains three oesophageal glands (two subventral, one dorsal) that overlap the nematode's intestine on the ventral side of its body. The intestine can be recognised as a fairly long dark region extending from the oesophageal gland cells to the tail. Secretory gland cells in the nematode oesophagus are the principal sources of secretions involved in plant parasitism (Hussey et al., 2002). The most evolutionary advanced adaptations for plant parasitism by nematodes are the products of parasitism genes expressed in their oesophageal gland cells and secreted through their stylet into host tissue (Davis et al., 2000). The secretions and putative effectors, both in sedentary and migratory endoparasitic nematode parasitism, mainly originate from the dorsal oesophageal gland cell and the subventral gland cells (Jones & Fosu-Nyarko 2014). For RKNs and CNs the subventral gland cells are the most active in infective second-stage juveniles, but following the onset of parasitism, the dorsal gland cell is stimulated to increase synthesis of secretory proteins to become the predominate gland later stages (Hussey & Mims 1990). The changes in the oesophageal gland cells during the parasitic cycle reflect changing roles of secreted proteins during different stages of parasitism.

The adult female can be recognised by the presence of the vulva (Fig 1.3), an opening in the cuticle on the ventral side, normally about 70-85% of the body length down from the 6

head of the nematode. Males are generally slightly smaller and more slender than females.

The defining characteristic of males is the presence of spicules (Fig 1.3); these are always present at the opening of the testis near the tail. Males have a row of cells that form

Figure 1.3. Adult female of P. curvicauda (see Chapter 2) (left) and a male of P. penetrans (right). the testis and the tail of males is more pointed that of females, and it often has two flaps of Figure 1.3. Adult female of ‘P. quasitereoides’ (left) and male of P. penetrans (right). cuticle attached. However, with so many proposed species, there are some morphological and morphometric characters which overlap, and so species identification can be difficult. For example, P. penetrans is closely related to P. fallax and differs only in having a crenate tail tip Figure(Seinhorst 1.3. Adult1968). female Several of characters ‘P. quasitereoides’ can be used (left) to identify and male nematode of P. penetrans species (right)in this . genus: the number of lip annuli, presence or absence of spermatheca in females, presence or absence of males, the number of lines in the lateral field and the shape of tail (Loof 1991;

Handoo & Golden 1989). Classically RLN species have been separated based on

morphometric parameters, but this is not always an exact process, because the values of different parameters can overlap, sometimes within a common range (Table 1.1). This means in practice that it is a challenge to differentiate some species based on morphology alone.

There can also be variation in morphometric parameters within the same species, depending on host, feeding, growth media, and geographically distant populations (Rashid & Khan

1976; Doucet et al., 2001). The body length and other morphometric values of RLNs can also be influenced by the state of the specimen being examined, in particular, whether live or fixed specimens are examined (Inserra et al., 2007). As a result, specific skills and longer term practical expertise is often required to identify RLNs based on their morphology and morphometry (Castillo & Vovlas 2007). Again, given the large number of species of RLNs, often little is known about intraspecific variability of morphological traits used for species identification (Tarte & Mai 1976), and it can be difficult to differentiate RLN species based on measurements of a single individual in a mixed RLN population. Thus, though the morphology provides significant information, molecular diagnostic tests, based on biochemical, molecular and phylogenetic analyses have emerged as powerful new tools to help refining the morphological definition of RLN species.

Table 1.1. Comparative morphometric measurements of some Pratylenchus species

P. quasitereoides P. teres P. curvicauda P. neglectus P. penetrans P. thornei P. zeae (Hodda et al., (Carta et al., (Siddiqi et al., (Mizukubo & (Ryss 1988) (Pourjam et al., (Van den Berg & 2014) 2002) 1991) Minagawa 1991) 1999a) Queneherve 2000) Body length (µm) 569-741 500-640 450-550 390-440 410-700 420-680 400-550 a* 14-25 20.0-9.8 21-28 19.8-26.0 19-30 23-37 21-31 (18) (24) (24) (23.1) (24) (31) (25.7) b* 6.3-8.7 3.7-4.9 5.2-6.8 5.0-6.2 5.3-6.7 5.2-8.0 4.5-6.5 (7.3) (4.4) (5.9) (5.5) (5.6) (6.5) (5.3) b’* 4.9-10.1 - 3-3.9 3.3-4.4 - 3.3-5.4 (7.0) (3.4) (3.8) (4.4) c* 15.9-22.5 15.4-8.4 13-18 16.6-20.3 15-24 15-26 13-19 (18.8) (16.8) (14.5) (18.1) (23) (20) (17.5) c'* 2.0-3.1 1.9-2.6 2.1-2.9 2.0-2.5 1.5-2.5 1.9-3.5 2.0-4.0 (2.3) (2.2) (2.7) (2.3) (2.1) (2.6) (2.5) V* (%) 75-82 71-77 69-76.5 80-82 77-83 72-82 74-80 (78) (75) (73) (81) (80) (77) (78)

*a=body length/greatest body diameter; b=body length/distance from anterior to oesophagus-intestinal valve; b'=body length/distance from anterior to base of oesophageal glands; c=body length/tail length; c'=tail length/tail diameter at anus or cloaca; V=% distance of vulva from the anterior end of the nematode body; values in parentheses represent average.

1.4 Molecular diagnostics and genetic diversity in RLNs

Molecular diagnostic techniques are mainly based on detection of specific nucleic acid sequences or on protein profiling, and molecular techniques are now used routinely to identify Pratylenchus species (Yan et al., 2012; Carrasco-Ballesteros et al. ,2007; Al-Banna et al., 2004; Subbotin et al., 2008; Tan 2012). Protein polymorphism and genetic diversity have been studied using Iso-Electric Focusing (IEF), such as for P. scribneri and P. brachyurus (Payan & Dickson 1990). Patterns of extracted proteins of Pratylenchus have also been analysed by sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS PAGE) for species identification, including: P. vulnus, P. goodeyi, P. coffeae, P. scribneri and P. thornei (Jaumot et al., 1997). Species-specific enzyme markers have enabled researchers not only to identify species but also to study intra-specific variation in populations from different geographical locations (Andrés et al., 2000).

Most protein- or isoenzyme-based diagnostic techniques have been superseded by

DNA and PCR-based techniques. These include RAPD, RFLP, ITS-RFLP, multiplex PCR,

SCAR-PCR, satellite DNA sequences (Yan et al., 2012; Carrasco-Ballesteros et al., 2007;

Al-Banna et al., 2004; Troccoli et al., 2008; Subbotin et al., 2008): their use has revolutionised molecular identification of Pratylenchus species, and provided invaluable data for phylogenetic analyses. Research on diagnosis of Pratylenchus in Australia now also makes use of these techniques, and nucleic acid based diagnostics (e.g. ITS based PCR) is used routinely to identify PPNs including species of RLNs (Tan 2012).

In parallel with nucleic acid based diagnostics, with advances in mass spectroscopy, it is also possible to use more recent rapid, protein separation methods, in particular matrix assisted laser desorption ionisation mass spectrometry time of flight mass spectrometry

(MALDI TOF MS) to identify nematode species of local and biosecurity concern, including

the RLNs P. neglectus, P. penetrans, P. thornei and P. zeae (Tan 2012). In addition, multiplex anti primer denaturation PCR (MAD PCR), which combines aQPCR technology and auto sticky PCR can increase the potential of detecting several species at the same time

(multiplexing) (Tan 2012). However, it is still important to combine new diagnostic technologies with morphometric diagnostics to develop accurate, reliable and rapid identification and quantification of known and emerging RLN species. This is because the accuracy of DNA based diagnostics is only as good as the initial identification of a species for which DNA sequence data is reported in sequence databases, and the initial diagnosis relies on experience and morphometric measurements.

1.5 Culturing, extraction and quantification of RLNs

Monoxenic culture of RLNs is useful for studying a particular species, from its identification to measuring the damage it causes. The first monoxenic culture technique was developed by Mountain (1960), who cultured P. penetrans on peach and tobacco tissues in vitro. Although various methods have been used for inoculum production of RLNs, the most widely used system is that of multiplying numbers on sterile carrot pieces, and this has been used successfully for maintaining and multiplying P. neglectus, P. thornei (Castillo et al.,

1995b), P. penetrans, P. brachyurus (Moody et al., 1973), P. zeae (Tan et al., 2013) and P. vulnus (Townson & Lear 1982a).

Because RLNs are migratory endoparasites, extraction of nematodes, from both living plant roots and the rhizosphere soil, including bulk soils, is necessary to quantify and determine their numbers. The Baermann funnel method is often used to extract nematodes from soil or roots, and extracts motile nematodes (Walker & Wilson 1960; Thorne 1961;

Hooper 1990). This process involves placing a funnel, with screen materials or fine cloth in its mouth, into a rack holder with a clamped rubber tube attached to the funnel. Soil samples, which can be wrapped in tissue paper, are placed on the screen material. Water is added to 11

the funnel just covering the soil sample and left overnight or longer. Nematodes slowly move through the screen and down the tube, and can be collected by releasing the clamp. The centrifugal flotation method is another technique for extraction of nematodes (Byrd et al.,

1966; Saunders & All 1982), which requires sucrose of high osmolarity and viscosity leading to long centrifugation times (Smith et al., 1997). In addition, use of iodixanol has also been effective in separating clean eggs and vermiform nematodes of Rotylenchus spp. (Deng et al.,

2008). The Whitehead tray method, with modifications (Whitehead & Hemming 1965;

Southey 1986; Doyle et al., 1987; Vanstone et al., 1998; Taylor et al., 2000) is most often used to extract nematodes from soil and roots. However, this method is effective only when inoculated plants or soil is moistened with water, and only live nematodes are extracted. For

RLNs, the mist technique, a modified Baermann funnel, and Whitehead tray are the most commonly used methods to extract nematodes from soil or roots. In this technique, soil or roots are put in a basket double lined with coffee filter paper placed on a funnel in a mist chamber, which provides a 10 seconds (s) water misting every 10 minutes (min). Active nematodes can be collected from a tube connected to the funnel.

There is a significant correlation between the population density of Pratylenchus in soil and extent of host damage (Ohbayashi 1989). The information on the number of RLNs present in a field helps in assessing future crop damage in that field. However, there is less published work on quantifying plant nematodes than, for example, soil bacteria or fungi

(Berry et al., 2008). Quantification of nematodes from soil and roots is done routinely by extracting live nematodes from soil or roots, followed by counting them under a dissecting or compound microscope. Collected nematodes can be separated based on their size and shape, through sieving, decanting or floatation. In contrast, DNA detection methods can now be applied as an effective and quick option to quantify nematode population densities in soil or roots. In this case quantitative real-time PCR is used to determine the nematode population

density based on the target DNA present and on the number of cycles of amplification required to first detect the nematode DNA (Yan et al., 2012; Nakhla et al., 2010).

1.6 Edaphic factors which affect RLNs

Soil texture, moisture, temperature and pH are the most important edaphic factors influencing survival, distribution and multiplication of RLNs (Castillo & Vovlas 2007). Soil texture is one of the important factors influencing the distribution of Pratylenchus species

(Castillo & Volvas 2007). For instances, most P. thornei were found in heavier textured soils

(48% clay, 37% sand) compare to P. neglectus (39% clay, 45% sand). The northern grain region of Australia appears to be more favourable for P. thornei than for P. neglectus

(Thompson et al., 2010). RLNs are adapted to a wide range of soil pH which correlates with specific host crops (Castillo & Vovlas 2007). Soil pH range from 6.5-9.5 are suitable for

RLNs. However, P. neglectus showed maximal incidence at 8.2 and P. thornei at 8.4. Soil temperature and moisture also affect reproduction, invasion and survival of RLNs (Kimpinski et al., 1976; Townshend 1973). For instance, although P. penetrans is a temperate species, low temperatures of about 5oC or less may be lethal, as found when potatoes were growing at this temperature (Olthof & Wolynetz 1991). Soil moisture regulates the vertical migration of

Pratylenchus species, and this is an important factor for diagnostic and faunistic studies.

Usually, Pratylenchus species are found from 0-105 cm in soil, with high variability depending on soil moisture, temperature, root distribution, temperature and rainfall (Castillo

& Vovlas 2007). Research on P. brachyurus on corn showed that the highest population was in the upper 30 cm of soil in summer and 20-40 cm in winter (Koen 1967), while in South

Australia, P. neglectus and P. thornei were mainly found in the top 20 cm of soil (Taylor &

Evans 1998).

1.7 Biology of RLNs

Pratylenchus species are polyphagous, obligate, migratory root endoparasites, although they may also behave as ectoparasites by feeding on root epidermal cells or root hairs without entering root tissues (Zunke 1990). Once inside the root, feeding is usually restricted to cells of the root cortex (Castillo & Vovlas, 2007). Unlike sedentary endoparasites, they do not develop any long term feeding sites in roots. The eggs are deposited singly or in clusters inside root cavities or in soil near the root surface (Acedo &

Rhode 1971; Bridge & Starr 2007; Castillo & Vovlas 2007; Pudasaini et al., 2008). P. neglectus, P. thornei and P. quasitereoides are parthenogenic with males being rare or absent

(Smiley & Nicol 2009; Carta et al., 2002). Conversely, P. penetrans is amphimictic having both males and females in the population (Smiley & Nicol 2009).

The life cycle (Fig 1.4) has four juvenile stages between the egg and adult (Embryo-

J1-J2-J3-J4-adult) (Luc et al., 2005). All the juveniles and adult stages are parasitic and can both enter and leave host roots. Information on the length of Pratylenchus life cycles under field conditions is limited: most studies on this aspect have been done in controlled conditions (laboratory and glass house). The life cycle of PPNs varies with nematode-host combinations, host, temperature and moisture (Loof 1991; Castillo & Vovlas 2007). RLNs can complete 3-6 generations per growing season (Taylor et al., 2000). Their life cycle is usually 24-65 days (d) depending on species and environmental variables (Smiley & Nicol

2009), as is indicated in Table 1.2. They can survive adverse conditions through an inactive, dehydrated state known as anhydrobiosis (Glazer & Orion 1983; Smiley & Nicol 2009). In this state, RLNs coil tightly to reduce the amount of exposed surface area and to retain water in summer (Glazer & Orion 1983; Tsai 2008) and also undergo diapause.

Figure 1.4. Life cycle of RLNs (Jones & Fosu-Nyarko 2014).

Table 1.2. Examples of the effects of host species and temperature on completion of RLN life cycles Figure 1.4. Life cycle of RLNs (Jones & Fosu-Nyarko 2014)

Pratylenchus Host Temperature Days to References spp. requirement complete life (°C) cycle P. coffeaeFigure Carrot1.4. Life callus cycle of RLNs (Jones30 & Fosu-Nyarko27-28 2014) . Wu et al., 2002 P. goodeyi Banana (resistant) 24 30 Prasad et al.,1999 Banana (susceptible) 24 24 P. loosi Carrot callus 20 45-46 Wu et al., 2002 P. mulchandi Pearl millet 25-30 24-36 Nandakumar & Khera 1974 Figure 1.4. Life cycle of RLNs (Jones & Fosu-Nyarko 2014) P. penetrans Red clover 17 46 Mizukubo & Adachi 1997 20 38 25 28 27 26 30 22 P. penetrans Carrot callus 24 34-35 Wu et al., 2002 P. thornei Carrot disc 20-25 25-35 Castillo et al., 1995a Corn 30 25-29 Siyanand et al., 1982 P. vulnus Carrot 26 26 Chitimbar & Raski 1985

1.8 Host range and pathogenicity of RLNs

Pratylenchus species have a broad host range, and this makes it difficult to find non- host crops for rotations to decrease populations below damaging levels. However, there is considerable diversity among Pratylenchus species in terms of pathogenicity to different hosts

(Vanstone et al., 2009). Individual crops, and even crop cultivars with resistance or tolerance to one species are not necessarily resistant or tolerant to another species of Pratylenchus

(Smiley 2009). For instance, wheat, barley and oats were all good hosts for P. neglectus

(Vanstone et al., 2009) and P. quasitereoides, while, canola can also be a good host for P. neglectus and P quasitereoides (Collins et al., 2017). Conversely, field pea, lupin and faba bean are poor hosts for P. neglectus and P. quasitereoides, but good for P. penetrans. The population density of an RLN species can also be influenced by different cultivars of the same host. For instance, cv. of canola, wheat and barley are good hosts for P. neglectus, but in barley, the multiplication rate of P. neglectus was greater on cv. Harrington than other cultivars

(Wilkinson et al., 2014, unpublished data). Similarly, the multiplication rate of P. quasitereoides was higher on the barley cv. Capstan than cv. Hindmarsh (Wilkinson et al.,

2014, unpublished data). Hence, cultivation of resistant cultivars or non-host crops can help to reduce the initial inoculum level and minimise the host damage in subsequent crops in a rotation.

Plants may be susceptible, resistant, tolerant or intolerant to nematode infestation.

Resistance is expressed as the ability of a host to inhibit nematode multiplication (Cook &

Evans 1987). Ideally resistance should be combined with tolerance, which is the ability of a

host to maintain yield potential in presence of nematode (Cook & Evans 1987). The actual

losses they cause for a crop, therefore, depend on initial population densities, crop plant and

status of cultivars used, soil conditions, rainfall and climatic factors. The pathogenicity of

Pratylenchus is influenced by soil temperature, texture or moisture, each of which in turn

affects nematode population densities (Wallace 1983). Added to these are variations in the populations of a particular nematode species that is variations in virulence. All of these factors need to be understood to comprehend the overall disease reaction (Shaner et al.,

1992). For example, dry shoot and root weights were influenced by the initial soil inoculum level and temperature for P. neglectus on alfalfa, where nematode reproductive indices were higher at 1000 nematodes per plant than 5000 per plant (Griffin & Gray 1990). The potential for crop damage is usually assessed based on initial inoculum of nematodes when planting

(Castillo & Vovlas 2007), and this number is important when deciding what control procedures to apply. Unfortunately for many species of RLNs, such information is often not available.

1.9 Host location and feeding of RLNs

The feeding behaviour of migratory endoparasitic nematodes has been less well documented than for sedentary nematodes (RKNs and CNs). In such studies, Pratylenchus nematodes have been described as movig through the root hair zone and using their lips and stylet probing, they rub the surface of selected epidermal cells (Kurppa & Vrain 1985; Zunke

1990). The invasion processes involves: recognition of the root, probing, penetration, ectoparasitic feeding, root entry and feeding from cells in the cortex, followed by moulting and reproduction (Linsell et al., 2014). RLNs sometimes show different responses or movements in terms of penetrating the roots. P. penetrans J2s were attracted to the root hair region in strawberry within 3 h of inoculation and penetrated through root hairs forcing their way into epidermal cells (Kurppa & Vrain 1985). In addition, a distinct coiling state was seen in P. penetrans just after entering a host cell, which was termed a rest phase, between feeding and further migration (Zunke 1990). However, P. thornei did not show different response in terms of penetration site in chickpeas root (Castillo et al., 1998), and usually migrated through epidermal and cortical cells by piercing cell walls along the migration pathway. It is

of interest to explore the changes that occur in host tissues at a cellular level in infected susceptible and resistant roots during Pratylenchus penetration, migration and feeding on root cells.

1.10 Histopathology and ultra structural changes in host cells infested with RLNs

Limited research has been carried out on the changes which occur in root tissues infested by RLNs. This is in contrast to the much more detailed ultrastructural studies which have been done by scanning or transmission electron microscopy (SEM or TEM) on the development and ultrastructure giant cells and syncytia in roots infected by RKNs and CNs roots (Jones & Dropkin 1976; Jones 1981). Most of the studies on histological or structural alterations in host cells have focused on P. penetrans. The biology of P. penetrans has been studied in tobacco, potato, oil radish and rape, and strawberry emphasising nematode penetration, migration to a feeding site, feeding, egg laying and moulting using high- resolution video-enhanced contrast microscopy and TEM (Zunke 1990). Histological responses of chickpea genotypes to P. thornei showed that damage from nematode infestation occurred from the epidermis inward to endodermal cells adjacent to the nematode feeding site

(Castillo et al., 1998). Changes at the cellular level have been studied in root cells of lily inoculated with P. penetrans (Vieira et al., 2017). However, more research is needed to detail the changes occur in infested roots of resistant or susceptible hosts.

1.11 Molecular studies of PPN-plant interactions

Although not the focus of this thesis, there have been rapid advances in molecular approaches to study events which underlie nematode-plant interactions, with most research focussed on RKNs and CNs. Among the many advances are understanding of differential activation of cell cycle gene promoters on infection with CNs or RKNs in Arabidopsis

(Goverse et al., 2000). Gene expression during the early stages of giant cells induced by

RKNs, have been studied by techniques such as laser capture microdissection (LCM), a powerful tool to study molecular changes directly in feeding cells (Ramsay et al., 2004), and use of FITC (fluorescein isothiocyanate) to tag live CNs (Schroeder & MacGuidwin 2007). In addition, behavioural studies of RKNs have been improved using Pluronic gel (Wang et al.,

2009). Laser scanning confocal microscopy provides a powerful strategy for studying early host-pest biological events associated with successful establishment infection stages of

Meloidogyne javanica and P. penetrans in A. thaliana using vital staining with FDA

(fluorescein diacetate) (Goto et al., 2010). There are a number of recent reviews that describe molecular advances in PPN-plant interactions (e.g. Gheysen & Mitchum 2011), and in particular for RLN-plant interactions (Jones & Fosu-Nyarko 2014; Fosu-Nyarko & Jones

2016), transcriptomes of P. zeae (Fosu-Nyarko et al., 2015) and P. coffeae (Haegeman et al.,

2011) and the first genome sequence, that of P. coffeae (Burke et al., 2015). The latter reports that P. coffeae has a remarkably small genome (about 20 megabases): further RLN genomes need to be sequenced to determine whether this figure is typical for other RLNs (Fosu-

Nyarko & Jones 2016).

One major area of advance is in the identification of a range of effectors, which are compounds secreted by PPNs which are needed for nematodes to be successful plant pests.

These include cell wall modifying enzymes (e.g. cellulases, pectinases), secreted effectors which avoid or evade plant defences, and proteins or peptides which modify plant cells by interacting with host components. The current status of effectors for Pratylenchus species is summarised in Fosu-Nyarko & Jones (2016): those present in both RLNs and sedentary endoparasites are core effectors, whereas those only present in RKNs or CNs are likely to be required for formation of giant cells or syncytia.

It is also clear that RLNs are amenable to host-induced gene silencing (HIGS), in which gene silencing (RNAi) technology can be used to silence vital genes, and so confer host resistance (Tan et al., 2013; Fosu-Nyarko & Jones 2016).

1.12 Conclusion

RLN infestation is particularly important in moisture-limiting soils, particularly towards the end of the growing season or in dry periods. It is of particular importance in

Australia, with rainfed cereals grown on low nutrient sandy soils, combined with no till agriculture to preserve topsoil. The predicted changes in global climate, particularly in south- western WA, will make this agricultural pathosystem even more prone to nematode damage.

Control of root diseases will therefore be critical to reduce economic losses. Although in these environments RLNs are economically the most important nematode pests, much less research has been done for these nematodes in terms of diagnosis, biology and nematode- plant interactions compared to sedentary nematodes. Most research has been directed to surveys of species present, pot and marker assisted screening of cereal cultivars for resistance and potential yield losses. There have been few studies using PredictaB to detect and quantify

DNA of specific species of RLNs in soil, but the issues of mixed species in the same soil or understanding the biology of particular species has yet to be studied in detail.

1.13 Aims and Objectives

The original impetus for the research described in this thesis was, P. quasitereoides

(formerly identified as P. teres), a relatively recently described nematode unique to WA

(Hodda et al., 2014), to fully characterise this speceis biologically and molecularly. However, it became clear early in this project that the sites selected for investigation were actually infested with P. curvicauda (evidence for this is provided in Chapters 2 and 3). This was an unexpected but highly fortuitous finding, because P. curvicauda has not been detected since

its original description from WA in 1991 (Siddiqi et al., 1991). Therefore, its biology, molecular biology and pest status are completely unknown, so to investigate these are of great importance for the cropping industry of WA. Consequently, the aims of the project were reframed as follows.

1) To undertake morphological and molecular characterisation of P. curvicauda in WA.

2) To study the genetic relationship of P. curvicauda with P. neglectus, P. penetrans, P. thornei and P. quasitereoides.

3) To study P. curvicauda life cycle, biology, penetration and attraction to resistant and susceptible hosts.

4) To study the host range of P. curvicauda.

5) To study histological and ultrastructural changes in susceptible and resistant host roots infested with P. curvicauda.

Chapter 2. Morphological characterisation of an undescribed Pratylenchus species from Western Australia

2.1 Introduction

RLNs are one of the most important biotic factors in Australia which limit grain yield, and they cause greater yield loss when plants are moisture stressed. For instance, P. thornei reduced water uptake by wheat roots in two years one with a wet and the other a dry growing season and that an intolerant cultivar lost 34% yield compared with a tolerant cultivar (Whish et al., 2014). P. neglectus and P. thornei are the two most common and economically important pest species for grains in different Australian states. Other species, such as P. zeae, are major pests of sugarcane production in finer textured soils in north-eastern Australia

(Blair & Stirling 2007). The extent of diversity of Pratylenchus species in WA is largely unknown (Riley & Wouts 2001). The species present in the WA grainbelt were thought to be predominantly P. neglectus (48%); other species present were P. quasitereoides (previously identified as P. teres), P. penetrans (1%), P. thornei (0.4%), and a mixture of species estimated to be as high as 26% (Collins et al., 2017). In this context, successful management could be undermined by a shift to predominance of other Pratylenchus species for which the crops grown were not resistant. This emphasises the need for accurate identification of the species present for successful management of RLN infestations. During the research undertaken in this project, soil samples were collected from four different locations of WA.

The initial morphological features were found as different from all the other four described species of Pratylenchus in WA. Therefore, the nematode samples were studied further to identify the species present in those field soils using morphometrics, light and scanning electron microscopy.

2.2 Materials and methodology

2.2.1 Extraction and culturing of nematodes

The nematode samples tested were collected from four different sites in the WA wheatbelt. The soil from Pingelly, Williams and Arthur River was collected by Sean Kelly,

Nematology Section, DAFWA. The sample from Katanning was provided by Helen Hunter,

Nematology Section, DAFWA in water after extraction. Barley cv. Hamelin was planted in pots containing the soils in a glasshouse to culture the nematodes that were present. After the plants had been growing for 8 weeks, nematodes were extracted from the soil using a misting apparatus as described by (Tan et al., 2013; Vanstone et al., 2008). This releases a fine mist of water for 10 s every 10 min. Live nematodes which had migrated through a double layer of coffee filters were collected after 72 h of mist treatment, and were used for morphological examinations.

To explore the relatedness of Pratylenchus species present in WA in terms of morphological features and genetic variation, three other known species of Pratylenchus, namely P. neglectus, P. penetrans and P. thornei, were also studied. These three nematode species were cultured on large carrot discs in vitro, and were originally provided as cultures by Dr. Vivien Vanstone, Nematology Section, DAFWA, having been started from individual identified nematodes. For culturing RLNs on carrot discs, healthy, good quality carrots were peeled using a sterile knife in a laminar flow cabinet. Both ends of the carrot were cut off, leaving a carrot piece of about 7 cm long. These were soaked in bleach solution (2.5%

NaOCl) with two drops of Tween 20 in a beaker for 30 min. The carrots were then peeled again to remove all bleached tissues and transferred to another beaker to soak them in distilled water. They were then cut into pieces (3-4 cm) to fit them into a sterile tissue culture jar, and the pieces then flamed after dipping in 70% ethanol before placing into the jar, which was left open in laminar flow cabinet for another 10-15 min to evaporate extra moisture from 23

the carrot pieces. After about a week, using a sterile scalpel, a small piece of older carrot cultured with nematodes was placed onto each carrot piece, and the jar at 20°C for culturing

P. neglectus and P. penetrans and at 25°C for culturing P. thornei.

2.2.2 Fixing nematode samples for measurement

Single individual nematodes from each sample were hand picked using a specially designed feather, and placed in a drop of water on a glass slide. They were then gently flamed, just enough to stop movement for study and taking photographs. The photographs were taken using a compound microscope (Olympus BX51). The morphological measurements viz. body length and the % distance of vulva from the anterior end of the nematode body, V were measured from the images using a scale bar.

Nematode specimens from Pingelly fixed in 4% formaldehyde solution were also sent to the UK, to Dr. M. R. Siddiqi, a renowned nematode taxonomist, for him to make detailed morphometric measurements.

2.2.3 Preparation of nematode samples for Light Microscopy

Live specimens of RLNs from the four locations and the other three known and closely related species (P. neglectus, P. penetrans and P. thornei) were hand picked using the feather, mounted in a drop of water on a glass slide and immobilised by gentle heating.

Photographs were taken using a Zeiss Axioskop Upright microscope (Carl Zeiss Microscopy,

LLC, United States) at the Centre for Microscopy, Characterisation and Analysis (CMCA) located at the University of Western Australia (UWA).

2.2.4 Preparation of nematode samples for Scanning Electron Microscopy (SEM)

A solution of 3% glutaraldehyde in 0.025 M phosphate buffer (pH 7.0) was used to fix adult female nematodes overnight at 4°C. The next day, the nematodes were washed 4-5 times in 0.025 M phosphate buffer. The specimens were post fixed using 1% osmium

tetroxide (OsO4) in 0.025 M phosphate buffer (pH 7.0) for 2 hours (h) at room temperature in a fume cupboard, followed by 4-5 washes in the same buffer. The samples were then dehydrated by washing sequentially with 30%, 50%, 70% and 90% ethanol twice in each solution, 15 min for each treatment. The 90% ethanol was then removed and replaced with

100% ethanol and then amyl acetate following the same regime. The specimen was then dried in a critical point dryer (FL-9496 BALZERS, Furstentum Liechtenstein). The nematodes were transferred to a SEM holder with conductive carbon tape and coated with a combination of 3 nm platinum and 10 nm carbon. Images were taken using an SEM (Zeiss Ultra 55) at 5

KV.

2.3 Results

2.3.1 Morphological features and measurements of nematode samples

As with most nematodes, the taxonomic designations of RLNs are based on the morphological characters of the adult female nematodes, because males are rarely present in populations of many species. The majority of nematodes from soils of the four locations were root lesion nematodes with typical, obvious stylets, with thick lip regions, presence of stylet knob, overlapping oesophageal glands, intestine at the ventral side of the body and a vulva located in the posterior third of the body (Fig 2.1).

Two parameters were measured initially: body length and value of V, % distance of vulva from anterior body of nematode. A total of 12 adult females were measured from

Pingelly, six from Williams, ten from Katanning and six from Arthur River (Table 2.1). The body length of the adult female from those locations varied from 441-768 μm with a mean of

538 μm, whereas V ranged from 70-76, with a mean of 74. The adult female from Pingelly was 484-616 μm (mean 550 μm) long with a V of 70-76 (mean 74); from Williams was 464-

590 μm (mean 520 μm) long with V value of 71-76 (mean 74); from Arthur River was 478-

636 μm (mean 534 μm) long with V of 70-76 (mean 73) and from Katanning was 441-768

μm (mean 538 μm) long with V of 72-77 (mean 74). No male nematodes were identified from the tested samples.

Figure 2.1. Morphology of the undescribed root lesion nematode from Pingelly; a. Adult female, b. head region, c. tail region (scale bar=100 μm).

Because Pratylenchus species have overlapping morphometric features, therefore, the obtained morphological measurements of this undescribed species were compared with those of RLN species known to be present in WA soils (P. neglectus, P. penetrans and P. thornei).

These measurements are presented in Table 2.1.

Table 2.1. Comparative morphometric measurements of Pratylenchus spp. adult female from WA

Pratylenchus spp. Pratylenchus sp. P. neglectus P. penetrans P. thornei Parameters (Undescribed) (WA) (WA) (WA) (n=34) (n=16) (n=16) (n=16) 441-768 403-552 447-616 557-809 Body length (538*) (480) (539) (650) (µm) 70-76 81-90 78-86 72-80 V (%) (74) (83) (83) (76) *values in parentheses represent the average values.

The measurements provided were the average values of 16 adult females from each of the three species. The body length of P. penetrans was 447-616 μm with a mean of 539 μm, which was within the range (410-700 μm) of previously published measurements (Ryss

1988). The body lengths of P. neglectus adult females was 403-552 μm (mean 480 μm) and that for P. thornei was 557-809 μm (mean 650 μm): these are slightly longer than previous measurements (Mizukubo & Minagawa 1991; Pourjam et al., 1999a), which may be because of geographical intraspecific variability (Corbett 1983; Castillo & Vovlas 2007). The value of

V ranged from 81-90 (mean 83) for P. neglectus, 78-86 (mean 83) for P. penetrans and 72-80

(mean 76) for P. thornei. Although the body length of these species overlapped, the position of vulva (%V) and tail shape indicates that the new nematode samples from the different WA locations clearly differ from those of P. neglectus, P. penetrans and P. thornei.

Although morphologically, using a compound microscope, this undescribed species looks like P. quasitereoides, as described from WA specimens by Hodda et al., (2014), however, based on morphometric measurements it did not appear to be as P. quasitereoides which had been described recently for WA. The average body length of the adult female of the undescribed specimens of Pratylenchus was shorter and the V value is less than that determined for P. quasitereoides (Hodda et al., 2014). This was a surprising result, since the initial aim of the research was to study the biology and molecular study of P. quasitereoides in more detail. Rather this appears to be an undescribed species of Pratylenchus. Hence, for additional detailed morphometric analyses, nematode specimens from Pingelly were sent to

Dr. M. R. Siddiqi.

2.3.2 Detailed description and morphometric analyses of nematode samples from Pingelly The root lesion nematodes extracted from Pingelly soils sent to Dr. M. R. Siddiqi for detailed morphometric measurements were described morphologically by him (Fig 2.2) as follows:

 The body of the nematode is irregularly curved to a c-shape; a maximum width is about 20 µm.  Annules are distinct, three in the head region.  Lateral fields with four incisures, the outer ones distinctly crenate, five to six incisures seen in vulval region.  The head is rounded, offset by a constriction, about 9 µm in diameter and 2.5-3 µm high with three distinct annules; framework strongly sclerotised, its outer margins extending two annules into the body.  The spear is strong, conus is 51-55 per cent of the spear length. The basal knobs are rounded: 5 µm across and 2 µm high.  The orifice of the dorsal oesophageal gland is at 3-4 µm behind basal knobs. The oesophageal glands are lying in tandem ventral to the intestine, whereas the dorsal gland is in anterior most.

 There is an anterior ovary with a single row of oocytes, not mature or reaching the oesophageal glands.  Sperm in spermatheca is not seen. The post vulval uterine sac is differentiated, with a few reduced cells of posterior ovary, 1.5-1.7 times vulval body width long.  The tail is sub cylindroid with terminal fourth usually appearing lozenge shaped, ventrally arcuate, terminus rounded, occasionally with an indentation.  Lateral field on tail with four incisures in anterior third, then with three incisures as it narrows down to end just before terminus.  Phasmid dot like, usually seven to nine annules or one fourth of tail length behind anal level. However, the longer conus (more than half of the spear length), three annules in lip region, large rounded basal knobs, phasmids in anterior region of tail, usually about one fourth tail length behind anus and the absence of males are good characters to compare the Pingelly specimens with related species.

The detailed morphometric measurements showed that the body length of the new nematode specimens from Pingelly was 415-540 µm with an average of 503 µm. The ratio of body length divided by greatest body diameter (a), was 20.7-27 with a mean of 24. The ratio of body length and distance from anterior to oesophago-intestinal valve (b) and the ratio of body length with distance from anterior to base of oesophageal glands (b ́) was 6-6.5 and 3.3-

3.37, respectively. The ratio of body length and tail length (c) was 12.9-14.6 and the ratio of tail length to tail diameter at anus or cloaca (c ́) was 2.4-3. The % distance of vulva from anterior body (V) ranged from 70-77, with a mean of 73.2.

Figure 2.2. Line drawings of different body parts of the undescribed Pratylenchus sample from Pingelly, WA: a-head, b-oesophageal region, c- vulva region, d & e-tail ends (as illustrated by Dr. M. R. Siddiqi, unpublished).

2.3.3 Comparative morphological features of the undescribed species from Pingelly and its relationship with other species of Pratylenchus

Based on these characters, the measurements seemed to be close to those of P. quasitereoides as described by Hodda et al., (2014). However, in the nematodes from

Pingelly, the lateral field is wider and has more incisures in the vulval region. The tail pattern is quite different since the inner incisures go past the phasmids. The phasmids of the Pingelly specimens are at the end of anterior fourth of the tail, while they are behind the middle of tail in P. quasitereoides: this feature is a very important morphological characteristic. Also, in the spear (stylet), conus is longer than the shaft including the basal knobs, and this appears to be larger in this undescribed specimens studied. Moreover, the tail of the Pingelly sample is 30

more arcuate ventrally and its terminus is smooth except for occasional single indentation, while P. quasitereoides was depicted as having a broadly rounded, finely crenate terminus tail.

A relative comparison was also made using the measurements with morphometric features of the same characters of other published Pratylenchus spp. including the species present or reported earlier in WA. This comparative study suggests that the nematode samples from Pingelly do not belong to any of the known species thought to be present in

Australia, and in particular in WA. The position of vulva and tail shape differs morphologically from those of P. neglectus and P. penetrans. For these two characterised species, the average value of V is 83, while for the undescribed specimen it is 74 (Table 2.1).

Similarly, P. thornei differs from the Pingelly samples and has a higher value of V (76). A similar trend in measurements was apparent from published data of P. neglectus, P. thornei and P. penetrans (Table 2.2).

Looking at the average value for ‘a’, the Pingelly samples matched to three species

(P. penetrans, P thornei or P. curvicauda) (Table 2.2). The values for ‘b’ or ‘c’ indicated that the new nematode samples could be from any of the six species. However, the value of b ́ for the Pingelly samples was 3.37 and for P. curvicauda it was 3.4, which were almost identical.

In contrast, this value was higher in other species compared in Table 2.2, and that reported for P. quasitereoides was almost twice the value measured for the undescribed Pingelly samples. Similarly, the value for c ́ for the Pingelly samples were 2.8 and was 2.7 for P. curvicauda, the value of c ́ ranged from 2.1-2.6 for P. neglectus, P. penetrans, P. thornei and

P. teres. In addition, the Pingelly specimens shared almost the same V value (73.2) only with

P. curvicauda (73), while for the remaining four species the value of V ranged from 75-81

(Table 2.2).

Table 2.2 Comparative morphometric measurements of undescribed Pratylenchus from Pingelly with other closely related species

Name of species Pratylenchus P. neglectus P. penetrans P. thornei P. curvicauda P. quasitereoides P. teres (Pingelly,WA) (Mizukubo & (Ryss 1988) (Pourjam et (Siddiqi et al., (Hodda et al., (Carta et al., Minagawa 1991) al., 1999a) 1991) 2014) 2002) Body length 415-540 390-440 410-700 420-680 450-550 569-741 500-640 (µm) a* 20.7-27 19.8-26.0 19-30 23-37 21-28 14-25 20.0-29.8 (24) (23.1) (24) (31) (24) (18) (24)

b* 6.0-6.5 5.0-6.2 5.3-6.7 5.2-8.0 5.2-6.8 6.3-8.7 3.7-4.9 (6.2) (5.5) (5.6) (6.5) (5.9) (7.3) (4.4) b ́ * 3.3-3.37 3.3-4.4 - 3.3-5.4 3-3.9 4.9-10.1 - (3.37) (3.8) (4.4) (3.4) (7.0) c* 12.9-14.6 16.6-20.3 15-24 15-26 13-18 15.9-22.5 15.4-18.4 Parameters (14.0) (18.1) (23) (20) (14.5) (18.8) (16.8) c ́ * 2.4-3.0 2.0-2.5 1.5-2.5 1.9-3.5 2.1-2.9 2.0-3.1 1.9-2.6 (2.8) (2.3) (2.1) (2.6) (2.7) (2.3) (2.2) V 70-77 80-82 77-83 72-82 69-76.5 75-82 71-77 (73.2) (81) (80) (77) (73) (78) (75)

*a= body length/greatest body diameter; b= body length/distance from anterior to oesophago-intestinal valve; b ́= body length/distance from anterior to base of oesophageal glands; c= body length/tail length; c ́= tail length/tail diameter at anus or cloaca; V= % distance of vulva from the anterior end of the nematode body; values in parentheses represent average values.

2.3.4 Features of nematode samples using Light and Scanning Electron microscopy

Light microscope images revealed that the undescribed specimens, this time present and studied from all four locations including Pingelly, differed from P. neglectus, P. penetrans and P. thornei, in terms of tail shape and position of the vulva from the anterior part of the nematode body (Fig 2.3).

With the conoid tail shape that is ventrally curved, terminus rounded with an indentation and the position of the vulva in a comparatively higher position, clearly separated the undescribed species of Pratylenchus can clearly be differentiated from P. neglectus, P. penetrans and P. thornei. The tail of P. penetrans is generally rounded, that of P. neglectus is conoid and for P. thornei the tail is conical with a round tip (Fig 2.3 & Fig 2.4). The tail of P. quasitereoides, however, is broadly rounded with a fine crenate terminus (Hodda et al.,

2014).

Figure 2.3 Differences in tail shape and relative position of the vulva of the undescribed sample labelled as Pratylenchus sp. and three other known species (P. penetrans, P. neglectus and P. thornei) examined using a compound microscope (moving nematodes). 33

Figure 2.4. Light micrographs of different species of Pratylenchus from WA. a. Entire female body (left), b. head (up) and c. tail region (bottom) of undescribed Pratylenchus from Pingelly; d. entire female body, e. head (up) and f. tail region (bottom) of P. neglectus; g. entire female body, h. head (up) and i. tail region (bottom) of P. penetrans; j. entire female body, k. head (up) and l. tail region (bottom) of P. thornei. (scale bar: a-c=50 µm and d-l=100 µm).

Using scanning electron microscopy (SEM), the front of the head or face was seen to be formed by the oral disc. In the samples from Pingelly the enface view was characterised by an oral disc which was slightly raised and divided. The head is rounded, offset by a constriction, about 9 µm in diameter and 2.5-3 µm high with three distinct annules (Fig 2.5 b

& c).

Figure 2.5. Female of undescribed Pratylenchus sp. from Pingelly under SEM. a. a whole nematode body, b-c. enface view, d. lateral field at vulval region.

The labial framework is strongly sclerotised, its outer margins extending two annuli into the body. The lateral fields have four lines, in which the lateral bands are wider than the

inner band. There was some irregular patterning in the mid body and tail regions, the outer ones distinctly crenate, with five to six incisures seen in the vulval region (Fig 2.5 d). The anus and vulva are clearly seen in the tail region (Fig 2.6 c & d). These morphological features were the same as described by Dr. M. R. Siddiqi for these nematode specimens.

Figure 2.6. Female of undescribed Pratylenchus sp. from Pingelly under SEM. a. tail

region, b. lateral field at middle of the body, c. tail terminus (arrow points to anus), d. vulval region (arrow points to vulva).

2.4 Discussion

The overall objective of this study was to characterise the RLN species collected from four different locations in the wheatbelt of WA with particular reference to the biology of P. quasitereoides, which had been described form WA by Hodda et al. (2014). The RLN samples provided for this project from DAFWA were thought to be P. quasitereoides, the initial results were puzzling, since the morphological and morphometric measurements did not fit the published parameters for this species. The taxonomic features used to separate

Pratylenchus at the species level are: the number of lip annuli, the presence or absence of a spermatheca in females, the presence or absence of males, the number of lines in the lateral field, and the shape of the tail (Handoo & Golden 1989). The identification of Pratylenchus species is usually based on female morphology since they possess more diagnostic characters than males if present (Loof 1991). Moreover, the female stylet length and % distance of the vulva from the anterior of the nematode are of high taxonomic significance, as these are less modified by biotic or abiotic factors (Castillo & Vovlas 2007).

The body measurements of the samples of the undescribed RLN specimens from the four WA locations demonstrated that they were morphologically different from P. neglectus,

P. penetrans and P. thornei. In addition, the measurements of samples of the undescribed

Pratylenchus did not match those of P. quasitereoides from Katanning, as described by

Hodda et al., (2014). The undescribed nematodes in the present study were from four locations including Katanning. The undescribed nematodes measured from each location were generally shorter than P. quasitereoides and had small V values. No type specimen of P. quasitereoides was available for comparative morphological studies.

The detailed morphometric measurements of nematodes from Pingelly and comparison with those of other species suggest that the undescribed specimens were not P. neglectus, P. penetrans or P. thornei or P. quasitereoides. The ratio of the body length and

greatest body diameter (a) were similar to those of P. penetrans, P. thornei or P. curvicauda, however, the other parameters such as the ratio of the body length and distance from anterior to oesophago-intestinal valve or the ratio of the body length and the tail length were not consistent with any of the six species compared. The value of the ratio of the body length and distance from anterior to base of oesophageal glands of the undescribed nematode specimens was in fact closer to that published for P. curvicauda, and this value was quite high in the case of P. quasitereoides. In addition, the ratio of the tail length and the tail diameter at the anus or cloaca in P. curvicauda and that for the undescribed nematode indicated the closeness between these two species. The other morphometric value of high taxonomic significance, the value of V in the undescribed specimens also pointed towards higher coincidence in morphology with P. curvicauda. In addition, the tail shape and phasmid indicated that the nematode samples studied in this project could well be P. curvicauda. In fact, this species had been reported earlier to be present in WA (Siddiqi et al., 1991). Analyses of the SEM images also supported the morphological description of nematode samples from Pingelly as P. curvicauda.

The findings in this study therefore suggest that the undescribed nematode examined in this study, including from Pingelly, is P. curvicauda.

2.5 Conclusion

This study confirms the importance of correct identification of RLNs and highlights aspects and difficulties in correctly identifying these nematodes at the species level. It also shows that the species of RLN collected from Pingelly was not P. neglectus, P. penetrans, P. thornei or P. quasitereoides. Rather, the morphometric measurements and the morphology suggest that it is very similar to P. curvicauda. It is important to resolve this taxonomic issue.

Its taxonomic validation will shed light on additional aspects of biology of this nematode. In the following chapters the species will be denoted as P. curvicauda.

Chapter 3. A study of genetic variation in Pratylenchus spp. in cereal growing regions of Western Australia

3.1 Introduction

RLNs, based on economic impact on crop yield, are one of the main concerns for cereal growers in WA. In the first published report involving P. quasitereoides, two or more RLN species were found together in 18% of surveyed paddocks, of which 13% had infestations of both P. neglectus and P. quasitereoides (Collins et al., 2013). In a more recent survey, of 380 paddocks in WA, 68% of infested paddocks showed the presence of single RLNs, while 32% had mixed RLNs (Collins et al., 2017). Current indications are that P. quasitereoides is the predominant RLN after P. neglectus in the wheatbelt areas of WA. Predicta B (B = broadacre) is a DNA-based soil testing service developed to identify and to assess risk of crop loss prior to seeding, which can minimise risk of yield loss. Data from PredictaB indicated that 56% of 340 paddocks tested were at risk of significant cereal yield losses

(Collins et al., 2017). The management of RLNs, either through crop rotations, cultural practices, nematicides or resistant crop varieties, always emphasises the accurate identification of the species present.

RLNs exhibit some overlapping morphological and morphometric characters which often make it difficult to distinguish between species. Within the same species, morphometric differences can also be present, depending on the host plant and geographic location, for example as found for P. penetrans (Townshend 1991; Tarte & Mai 1976). Using classical taxonomy, high-level skills and experience are required to identify RLNs (Castillo &Vovlas

2007). Because of this skills requirement, molecular approaches and phylogenetic analyses are being used increasingly as powerful tools for nematode diagnostics, offering greater accuracy and reproducibility, as long as the original specimens used for molecular diagnostics were identified correctly using traditional methods. As for many other organisms,

the genes encoding the rRNA subunits have proved to be useful in taxonomic studies of nematodes. These genomic regions vary in their rate of evolution depending on whether they encode functional products or not. These sequences include those of the ribosomal small subunit genes which can be extremely conserved, or the non-coding internal transcribed spacer (ITS) regions which are much more variable between species of the same genera

(McKeand 1998). Sequencing of the ITS regions has revealed species-specific variations, which can be used as diagnostic markers, so enabling accurate identification of species and studies on the phylogenetic relationships between and within species such as Pratylenchus

(Waeyenberge et al., 2009; Palomares-Rius et al., 2010). In addition, the nucleotide sequences of D2-D3 from the large subunit ribosomal genes (28S), which evolve slowly, can be used to examine the evolutionary relationships among species of the genus Pratylenchus

(Al-Banna et al., 1997).

Information on genetic variation between and within nematode species is important when considering breeding for nematode resistance. Work in this chapter is focused on P. curvicauda collected from four different wheatbelt regions of WA, and other known species of Pratylenchus (P. neglectus, P. quasitereoides, P. penetrans and P. thornei), with the aim of determining the genetic variation between species and in Pratylenchus populations based on both the ITS region and the D2-D3 expansion segments of the 28S rDNA gene.

3.2 Materials and methods

3.2.1 Nematode populations, extraction and culture

The work in this chapter involved study of the genetic variation of P. curvicauda with four Pratylenchus spp. identified in cereal fields of WA as P. neglectus, P. quasitereoides, P. penetrans and P. thornei. Nematode samples of P. curvicauda were obtained from four different locations in the wheatbelt region of WA: Pingelly (32°32′2.4″S, 117°5′9.6″E),

Williams (33°2′0″S and 116°53′0″E), Arthur River (33°20′19″S and 117°2′4″E) and

Katanning (33°41′27″S and 117°33′19″E) (Fig 3.1). Soils from these locations were thought to be infested with P. quasitereoides (Department of Food and Agriculture, Western

Australia, DAFWA, Dr. Sarah Collins).

Figure 3.1 The four locations from where Pratylenchus samples were collected.

Soil samples were first sieved using a 1 cm mesh and then mixed to homogeneity to obtain representative subsamples. Nematodes were extracted from soils using a misting apparatus as described by Tan et al., (2013). Soil samples (100 g) were placed on two coffee filters on a funnel over an open syringe with its outlet sealed by a clip on a silicon rubber tubing. Nematodes were extracted by release of a fine mist of water every 10 min for 10 s.

Nematodes were collected from the syringe by releasing the clip on the silicon tubing 72 h after the soil was placed in the misting apparatus, and used for further morphological and molecular work. P. neglectus, P. penetrans and P. thornei, which had been collected

previously from WA soils by DAFWA staff, were maintained on carrot disks at 21-25°C depending on the species. Primary identification of Pratylenchus species was carried out using two important morphological features: the position of the vulva and the shape of the tail. A number of single adult females of each species were examined under a compound microscope to identify species accurately before DNA extraction. For photographic recording, nematodes were immobilised by gentle flaming on a glass slide. All measurements of nematodes were done based on comparisons with micrometer (µm) scale bars, and the percent distance of vulva from anterior V was calculated.

3.2.2 DNA extraction

Single adult female nematodes were hand picked using a sharply pointed feather, placed in a 10 μl drop of sterile water on a glass slide and cut into two or more pieces under a dissecting microscope. Genomic DNA was extracted using a modified protocol involving a worm lysis buffer (Wood 1988), followed by phenol-chloroform extraction. Nematode fragments in 10 μl of water were transferred to a sterile tube with 50 μl of lysis solution (1%

SDS, 50 mM EDTA, 100 mM NaCl, 100 μg/ml proteinase K, 1% 2-mercaptoethanol, 100 mM Tris-HCl pH 8.5). The nematode lysates were frozen at -80◦C for 40 minutes followed by thawing and heating at 60◦C for 40 minutes. The supernatant was transferred to a fresh tube for subsequent extraction using an equal volume of phenol-chloroform (50:50). A one-tenth volume of sodium acetate (3M NaOAc, pH 6.8) and 100% ethanol (2.5 volume) were used to precipitate DNA at 16,000× g. The pellets were washed twice with 400 μl of ethanol (70%), dried in a fume hood and resuspended in 17μl of nuclease-free water. The DNA was quantified using a Spectrophotometer (Nanodrop ND-1000, Isogen Life Sciences) and stored at -20°C until use.

3.2.3 PCR amplification, cloning and sequencing

The partial 18S-ITS1–5.8S-ITS2-partial 28S regions of the rDNA fragment (Fig 3.2) were amplified by PCR using the primer pairs 18S-Int (5´-CGTAACAAGGTAGCTGTAGG-

3´) and 26S-Int (5´-CCTCCGCTAAATGATATGC-3´) (De Luca et al., 2011). The DNA was amplified using GoTaq® Green Master Mix (Promega Corporation, Australia), a premixed ready-to-use solution containing bacterially derived Taq DNA polymerase (50 units/ml,

400µM dATP, 400µM dGTP, 400µM dCTP, 400µM dTTP, 3mM MgCl2). The reaction conditions were: 95°C for 5 min; 35 cycles of 95°C for 30 s, 55°C for 5 s, 72°C for 1.40 min followed by a final step of 72°C for 10 min.

Figure 3.2. The location of the primers used to amplify different regions of rDNA genes of Pratylenchus spp.

Also the D2-D3 regions of the 28S gene was amplified using the primer pairs D2-F

(5’Figure-GACCCGTCTTGAAACACGGA 2.2. The location of the primers-3’ )used and D3to amplify-R (5’-TCGGAAGGAACCAGCTACTA different regions of rDNA genes -

3’of) (De Pratylenchus Luca et al., spp. 2004) (Fig 3.2) with the following temperature profiles: 94°C for 6 min, followed by 35 cycles of 94°C for 1 min, 55°C for 1 min, 72°C for 1 min and a final step of

72°C for 6 min. PCR products were run on 1% gel stained with SYBR Safe (Invitrogen Pty

Ltd, Australia). The respective bands from gels were cut out and purified using the Wizard®

SV Gel and PCR Clean Up System (Promega Corporation, Australia) and purified DNA was used for Sanger sequencing. Because direct sequencing of the amplicons yielded ambiguous peaks, the PCR products were subsequently ligated to the pGEM-T Easy vector (Promega

Corporation, Australia) and the plasmid used to transform JM 109 competent cells as described in the manufacturer’s protocols (Promega Corporation, Australia). For amplicons 43

from the ITS region, a total of six clones from each single nematode were selected and sequenced. For amplicons obtained from the D2-D3 region, one clone from each single nematode was sequenced. Plasmid DNA from cultures was purified using the

Wizard®PlusSV Minipreps DNA Purification System (Promega Corporation, Australia).

Both strands of DNA fragments from each clone were sequenced. Sequencing was done using an AB 3730 96 capillary DNA sequencer (Applied Biosystems).

3.2.4 Analyses of sequences

Raw sequences were visualised and edited with Finch TV (version 1.4.0) (Geospiza

Inc. 2004). The sequences were aligned using CLUSTAL O (1.2.3) (McWilliam et al., 2013) and Geneious (V 8.1.8) (Biomatters; Kearse et al., 2012) and the same programs were used to identify homology and conserved regions within sequences. BLAST searches of ITS sequences at NCBI (https://blast.ncbi.nlm.nih.gov/Blast.cgi) were used to retrieve similar sequences of Pratylenchus spp. The D2-D3 sequences generated in this project were also compared to the same region and length of similar sequences of Pratylenchus spp. available in NCBI databases.

Phylogenetic analyses were carried out using MEGA 6 (Tamura et al., 2013). The

Maximum Likelihood (ML) approach was chosen to construct the trees. The model used for generating phylogenetic trees was selected as the best fit among standard models for both the

ITS and D2-D3 dataset and models with the lowest BIC (Bayesian Information Criterion) scores were considered which described the substitution pattern best. Sequences from M. hapla and Radopholus similis were used as outgroup for phylogenetic trees of ITS sequences, whereas a sequence of Belonolaimus longicaudatus was used for D2-D3 sequences. In all cases, bootstrap values were based on 1000 replicates.

3.3 Results

3.3.1 Morphology of the Pratylenchus species studied

The majority of nematodes in soils from each of the four locations were plant parasitic nematodes because they had a characteristic mouth stylet. Close observation of individual nematodes indicated that they were RLNs because they had a thick lip, a stylet knob and overlapped oesophageal region. No male was observed during microscopic examination.

According to the morphometric measurements including the position of the vulva and the tail shape, the nematode specimens from the four locations, were clearly different from P. quasitereoides, P. neglectus, P. penetrans and P. thornei. All the details have already been described in the previous chapter (Chapter 2).

3.3.2 Length of ITS sequences of P. curvicauda, P. neglectus, P. penetrans and P. thornei

The primer pairs were used successfully to amplify the intended ITS regions of DNA from single adult female nematodes of all the Pratylenchus species used in this study. The amplicon size was approximately 900 bp on a 1% agarose gel (Fig 3.3). Attempts were made to sequence the amplicons directly, but it appeared that sometimes there were multiple products. DNA fragments were then sequenced after cloning. Based on the sequencing results, the lengths of the amplified ITS sequences of the clones varied.

Figure 3.3. Amplified ITS region of Pratylenchus spp.; 1. P. curvicauda, 2. P. neglectus, 3. P. penetrans, 4. P. thornei, 5. Negative control; M=100 bp marker.

Figure 2.3. a. Amplified ITS region of Pratylenchus spp.; 1. ‘P. quasitereoides’, 2. P. neglectus, 3. P. penetrans, 4. P. thornei, 5. negative control; b. Amplified D2-D3 gene segments of

All the ten clones of P. curvicauda from Williams and Arthur River were 906-907 nucleotides (nt) long: the amplified ITS regions were slightly longer for the clones from

Katanning (928-933 nt) and Pingelly (906-933 nt). Amplicons of the ITS regions from P. neglectus (825-826 bp) and P. thornei (826 bp) were shorter compared to sequences from the

P. curvicauda from the four locations, whereas 12 clones from the amplified ITS region for

P. penetrans was between 930-933 bp long. For all four RLN species, the length of the ITS1 sequence was between 311-392 nt and of the ITS2 was between 268-342 nt.

3.3.3 Variation in ITS sequences of P. curvicauda

Overall, 28 clones of amplicons for P. curvicauda from the four locations were sequenced and analysed. The ITS sequences covering the ITS1 and ITS2 regions of P. curvicauda showed some intra-specific heterogeneity. The sequences from Katanning showed 0-6% variation overall, while, the clones from Pingelly showed 0-9% variation.

Regarding the ITS1 region, sequences from Pingelly and Katanning showed 0-12% (0-46 nt) variation and in the ITS2 region the differences were 0-11% (0-36 nt). However, the clones from Williams and Arthur River were less variable (less than 1%). In particular, the ITS1 and

ITS2 regions in the sequences from Williams and Arthur River were almost identical (99-

100%). The sequences of the amplified ITS region, without the ITS1 and ITS2 (i.e. the partial

18S, 5.8S and partial 28S were conserved among the 28 clones with only 2-3 nucleotide differences.

3.3.4 Variation in ITS sequences between P. curvicauda, P. neglectus, P. penetrans and

P. thornei

All 28 clones of P. curvicauda from the four locations were compared with sequences of similar regions of the ITS sequences of P. neglectus (6 clones), P. penetrans (12 clones) and P. thornei (5 clones). Sequences of clones from each of P. neglectus, P. penetrans and P. thornei were 100% identical. However, P. curvicauda sequences were only 61% identical to

those of P. neglectus and P. thornei. Conversely, the ITS sequences of P. curvicauda from the four locations were between 90-99% identical to those of P. penetrans. Sequences of P. curvicauda clones collected from Katanning and three clones of the same species from

Pingelly (PN2) were more similar to those of P. penetrans with less than 1% difference, whereas the differences in nucleotides were up to 10% in sequences of all the P. curvicauda clones from Williams, Arthur River and three clones of the same nematode from Pingelly

(PN1). These results indicated that the nematodes from the same field may constitute a mixed infestation.

Variations between sequences of P. curvicauda from the four locations and those of

P. neglectus, P. penetrans and P. thornei were mostly confined to the ITS1 and ITS2 regions; there were 44% and 49% variation in the ITS1 and ITS2 with P. thornei and P. neglectus with only 10% in both regions compared to P. penetrans sequences. Similar differences were found in region-specific alignment of ITS sequences, and revealed that the sequences of P. curvicauda were genetically very different based on the sequences of ITS1 (44%) and ITS2

(49%) regions of P. neglectus and P. thornei, and were more similar (10%) to ITS1 and ITS2 of P. penetrans.

3.3.5 Comparison of ITS sequences of RLNs from WA and global data

When the sequences of RLNs from the WA wheatbelt were compared with similar data generated internationally, the ITS sequences from P. neglectus and P. penetrans from

WA were considerably different. P. neglectus and P. penetrans sequences differed by 36-

66% and 43-48% respectively, with the global populations of the same species. In contrast, P. thornei sequences matched well (96-99.9%) with corresponding sequences in the GeneBank database (NCBI), except for two sequences from Italy (FR692303) and Spain (FR692305).

Interestingly, for WA P. neglectus, there was only 62% similarity in the ITS1 and 36% in the

ITS2 with 20 sequences from Italy. Similarly, with seven sequences of P. penetrans from the

USA: there was 60% similarity for the ITS1 and 38% for the ITS2 with those from WA.

Although soil collected from the four locations were previously known to contain nematodes that had been identified as P. quasitereoides, in the work which described this species (Hodda et al., 2014) its ITS rDNA sequences had not been examined, and so there was no sequence information for ITS rDNA from P. quasitereoides with which to compare with the sequences obtained in this study. However, there was some sequence data for P. quasitereoides for the D2-D3 region.

3.3.6 Analyses of D2-D3 sequences

The D2-D3 sequences of P. curvicauda were therefore generated for the samples collected in this study. A total of 17 sequences derived from 17 individual adult females obtained from the same four locations in WA were analysed. Also 14 sequences from P. neglectus, P. penetrans and P. thornei were analysed and compared to those of P. curvicauda. Following

PCR of the D2-D3 regions, from agarose gel electrophoresis, the size of the PCR products was approximately 300 bp (Fig 3.4). Sanger sequencing yielded sequences of between 312 and 316 nt. When subdivided into the four regions, the amplified D2-D3 region of six

Figure 3.4. Amplified D2-D3 gene segments of Pratylenchus spp.; 1. P. curvicauda, 2. P. neglectus, 3. P. penetrans, 4. P. thornei, 5. Negative control, M=100 bp marker.

48 Figure 2.3. a. Amplified ITS region of Pratylenchus spp.; 1. ‘P. quasitereoides’, 2. P. neglectus, 3. P. penetrans, 4. P. thornei, 5. negative control; b. Amplified D2-D3 gene segments of Pratylenchus spp.; 1. ‘P. quasitereoides’, 2. P. neglectus, 3. P. penetrans, 4. P. thornei, 5. negative control, M=100 bp marker. nematodes from Pingelly (PN) were 312-316 nt long, while those of four sequences from

Williams (WL) were 314-316 nt, three sequences from Arthur River (AR) were 313-316 nt long and four sequences from Katanning (KAT) were all 318 nt long. In comparison, the sequenced D2-D3 region of six P. neglectus and five P. penetrans sequences were 314-316 nt in length, and those of three P. thornei nematodes were 345-346 nt. Analyses of the D2-D3 sequences of P. curvicauda indicated that there was some variation in sequence between nematodes from different locations. In particular, nucleotide sequences from Pingelly varied by up to 12%, and sequences of nematodes from Williams varied by up to 8% and those from

Arthur River varied up to 9%. Conversely, the sequences of nematodes from Katanning were more similar, with less than 1% nt variation. Three sequences from P. thornei and six sequences of P. neglectus exhibited up to 8% nt variation, whereas, five sequences of P. penetrans showed up to 18% variation. This clearly indicates that there is heterogeneity in the

D2-D3 sequences of Pratylenchus sp. as was the case for the ITS sequences. All the 17 sequences of P. curvicauda from the four locations, and 14 sequences from P. neglectus, P. penetrans and P. thornei, differed by up to 30%.

3.3.7 Comparison of D2-D3 sequences of P. curvicauda with those of P. quasitereoides

A total of 17 sequences generated from P. curvicauda collected from the four locations were aligned with ten P. quasitereoides and two P. teres sequences available in

NCBI. Sequence alignments were done using sequences of the same region with similar lengths. The 17 sequences obtained in this study were very different from those reported by

Hodda et al (2014), with up to 39% variation. There were clear indels in the bases at positions 61-62, 88-102 and 116-131 of the sequences as presented in Fig 3.5. The amplified

D2-D3 sequences of P. quasitereoides from Hodda et al. (2014) was 215 bp, therefore, they were compared with the same length sequences from 17 nematodes obtained in this study.

Figure 3.5. Multiple alignments of sections of D2-D3 with variation in 17 sequences of P. curvicauda and published D2-D3 sequences of P. quasitereoides: AR- P. curvicauda sequences from Arthur River, WL- P. curvicauda sequences from Williams, PN- P. curvicauda sequences from Pingelly, KAT- P. curvicauda sequences from Katanning and D2-D3 sequences of P.quasitereoides from Hodda et al. (2014), as marked with the accession number from KM248297.1-248306.1. The numbers denote the number or identity of clones from

the same location).

3.3.8 Comparison of D2-D3 sequences

Figure 2.4. Multiple alignments of D2-D3 sequences from the four locations of WA: AR-‘P.quasitereoides’ sequences fromThe Arthur D2 -River,D3 sequences WL-‘P.quasitereoides’ of P. neglectus sequences, P. penetrans from Williams and P. thornei, PN-‘P.quasitereoides’ obtained in this sequences from Pingelly, KAT-‘P.quasitereoides’ sequences from Katanning and D2-D3 sequences of study were compared with sequences of the same species and regions from other locations ‘P.quasitereoides’ from Hodda et al. (2014), as marked with the accession number from KM248297.1- 248306.1).around These the show world. the This region analysis where thedemonstrated differences inheterogeneity sequences (the in fullsequences length of from the sequence different is 215bp, only part of the sequences are shown here). locations. Six P. neglectus D2-D3 sequences were up to 30% different from those of the same

species from the USA, China and Iran. Similarly, five sequences of P. penetrans showed 1-

18% variation and comparison with seven sequences from Japan, Netherlands and Chile

showed up to 30% variation. Although the three P. thornei sequences generated in this study

were 98-100% similar, they differed by about 9% from eight sequences of P. thornei from

South Australia and Queensland.

3.3.9 Phylogenetic analyses of ITS and D2-D3 sequences

The phylogenetic tree constructed using the amplified ITS sequences of P. curvicauda from the four locations and P. neglectus, P. penetrans and P. thornei shows that all the clones from Katanning and three clones of one P. curvicauda nematode from Pingelly were grouped together with those of P. penetrans (cluster I- Fig 3.6). All clones from Arthur River,

Williams and three clones from Pingelly were in cluster II. Two individual nematodes sequenced from Pingelly were in a separate clade indicating genetic differences in individual nematodes collected from the same location. In cluster III, P. neglectus and P. thornei were grouped together. Also grouping of some of the P. curvicauda with P. penetrans indicates, for the region of the ITS analysed, some nematodes from the four locations, particularly

Katanning may be more similar to P. penetrans.

curvicauda sequencesfrom PN Williams, appropriateclades. Likelihood bas neglectus Fig

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Phylogenetic sequences relationships betweenITS of

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All the ITS sequences of P. curvicauda from the four locations and those of P. neglectus, P. penetrans and P. thornei were compared with similar regions of 156 sequences from 26 different

Pratylenchus species available at NCBI to study relatedness of these species. Maximum likelihood using the GTR method with the invariant site plus gamma options resulted in a tree where sequences of P. quasitereoides formed a highly supported separate clade in which P. bolivianus was the closest species followed by P. bhattii (Fig 3.7). There is not any ITS sequence data for P. teres. The four

ITS sequences of P. bolivianus used to construct the phylogenetic tree were from the United

Kingdom, Chile and China, whereas all three sequences of P. bhattii were from Hong kong (Fig 3.7).

The sequence details with accession number of the Pratylenchus spp. used for phylogenetic analyses are listed in Appendix 1.

The P. neglectus, P. penetrans and P. thornei ITS sequences in this study were longer than corresponding sequences of the same species from locations other than WA. The ITS sequences from

P. neglectus were clustered separately with corresponding sequences from Italy (Fig 3.8 a), and P. penetrans displayed similar cluster characteristics when compared with sequences of the same species from Belgium, Chile, France, Netherlands and USA (Fig 3.8 b). Conversely, P. thornei sequences formed a well-supported cluster, except two accessions namely FR692303 and FR692305 from Italy and Spain respectively (Fig 3.9).

number number detailsand of the ITS sequences are listed in App four locations of WA obtained in this study are labelled with black a dot. The accession Evolutionary analyses were conducted MEGA6. in The Figure 2.6 details of the ITS sequences locations of WA obtained this studyin are labelled black a with dot. The accession number and Phylogenetic Figure 3.7.

Maximum Likelihood (GTR+G) tree using ITS the sequences o

Maximum Likelihood (GTR+G) tree using ITS the sequences of

analyses were conducted MEGA6. in The

other other

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endix 1. endix

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Figure 3.8. a. Maximum Likelihood (K2+G+I) tree constructed from the ITS of P. neglectus WA population (Pneg1.1-Pneg3.3) with corresponding sequences from GeneBank. Evolutionary analyses were conducted in MEGA6; b. Molecular Phylogenetic analysis by Maximum Likelihood method (T92+G) showing relationships among global population of Pratylenchus penetrans. Ppen1.1-Ppen5.2 is P. penetrans form WA.

Figure 3.9. Molecular phylogenetic analysis of ITS sequences of P. thornei by Maximum Likelihood method (K2: Kimura 2-parameter), Pth1.1-Pth2.2 indicates P. thornei from WA.

Figure 2.8. Molecular phylogenetic analysis of P. thornei by Maximum Likelihood method (K2: Kimura 2-parameter), Pth1.1-Pth2.2 indicates P. thornei from WA.

The phylogenetic tree constructed using the D2-D3 sequences of P. curvicauda from the four locations with 12 available sequences of P. quasitereoides and P. teres from NCBI shows that the P. curvicauda sequences studied in this thesis were different from the sequences used in Hodda et al

(2014) and formed a well-supported clade (Fig 3.10). The D2-D3 sequences of P. curvicauda obtained in this study were compared with a total of 48 corresponding sequences of 21 different

Pratylenchus species from different geographic locations (Fig 3.11). The P. curvicauda sequences from the four locations of WA were placed in a separate clade, which indicates that the sequences are quite likely to be from a different species of Pratylenchus.

Figure 3.10. Phylogenetic relationships of P. curvicauda sequences from the four locations in WA and sequences of P. quasitereoides from Hodda et al., (2014) as inferred from D2-D3 sequence alignment under K2+G model. P. quasitereoides from Hodda et al., (2014), as marked with the accession number from KM248297.1-248306.1, named as Pratylenchus sp. Posterior probabilities more than 50% are given for appropriate clades. Newly obtained sequences of P. curvicauda from this study are indicated by a black dot.

sp. al.et locations of WA usingMaximum Likelihood K2+G model. Fig (2014), of WA usingMaximum Likelihood K2+G model. Figure 3.11

ure 2.10

(2014),

as as marked with the accession number from KM248297.1

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3.4 Discussion

This work appears to be the first study of genetic diversity within and between

Pratylenchus spp. in Australia based on both D2-D3 and ITS sequences. The overlapping morphological features, the presence of morphotypes, together with a higher level of intraspecific and interspecific genetic variation in Pratylenchus, complicate the identification and separation of these species. However, recent advances in DNA sequencing and phylogenetic analyses using both ITS and D2-D3 regions of rDNA have facilitated the assessment of heterogeneity among species including those which are closely related. The shorter length of the D2-D3 region facilitates its amplification and it has been suggested that this region is a better target for studying higher degrees of interspecific genetic variation

(Subbotin et al., 2008). However, the ITS region appears to evolve faster than the D2-D3 expansion region, and is also known to contain more substitutions (De Luca et al., 2011).

Hence, this variation in sequence is probably more useful for identification of nematodes at the species level, and its study has allowed the resolution of complex species issues in

Pratylenchus.

This study has shown that it can be difficult to separate Pratylenchus at the species level, and hence it is important to combine both morphology and molecular data in identifying species accurately. The distance of vulva from the anterior end of the nematode body and the stylet length of the female is of taxonomic importance and are reliable diagnostic characters in Pratylenchus, as these characters are less modified by biotic and abiotic factors (Castillo &Vovlas 2007). The RLNs collected from soils of Pingelly,

Williams, Arthur River and Katanning were thought to be P. quasitereoides based on the morphology, their tail shape and the relative position of the vulva (%V). Later the species was identified as P. curvicauda after detail morphometric analyses, the results of which

differed from those of P. neglectus, P. penetrans and P. thornei descibed in Australia. This difference was confirmed later by sequencing and analyses of the ITS and D2-D3 regions.

Both the amplified D2-D3 and the ITS regions of P. curvicauda from the four locations varied both in length and nucleotide sequences. Similar intraspecies and interspecies divergence has been reported for other RLNs (Waeyenberge et al., 2000, 2009;

De Luca et al., 2011). Intraspecies variation of ITS sequences of Pratylenchus was evident in

P. coffeae (Duncan et al.,1999; Waeyenberge et al., 2000) and P. parazeae (Wang et al.,

2015), in P. agilis, P. brachyurus and P. penetrans (Waeyenberge et al., 2009). The ITS sequences obtained in this study from different clones as P. curvicauda also showed sequence heterogeneity. This variation could be caused by the presence of multiple and highly divergent copies of the ITS region in the nematode, as suggested for Meloidogyne sp. (Hugall et al., 1999). Cloning of DNA from a single nematode, amplifying any possible ribosomal gene irrespective of abundance and functional character, resulted in some heterogeneity between clones. This heterogeneity may arise from slightly different rDNA sequences in individuals or the presence of pseudogenes (De Luca et al., 2011).

The ITS sequences of P. quasitereoides from the four locations clearly distinguished them from P. neglectus, P. penetrans and P. thornei. The primer pair used to amplify the ITS region of Pratylenchus spp. in the study has previously been used for several species

(Waeyenberge et al., 2000; De Luca et al., 2011). Interestingly, the length of amplicons obtained ranged from 900 to 1250 bp. For example, the length is 900 bp for P. neglectus from

Spain and is between 900 and 1000 bp for P. penetrans from Japan, whereas it is 1200 bp for

P. thornei from Spain (Waeyenberge et al., 2000). The same primers yielded amplicons ranging from 693 to 1170 bp for Pratylenchus spp. including P. neglectus and P. thornei (De

Luca et al., 2011). The ITS sequences of P. neglectus, P. penetrans and P. thornei obtained

in this study were longer compared with corresponding sequences of the same species available in GeneBank.

One result is that the P. curvicauda sequences were more similar to those of P. penetrans than to P. neglectus or P. thornei. P. penetrans was also the most variable of the

RLN population in WA. Also, the ITS region of P. neglectus and P. thornei differed from the corresponding sequences from the same species in other regions of the world. Sequence divergences were also found between P. parazeae. and P. zeae population from Taiwan with other locations (Wang et al., 2015). The results reported here also show that the ITS1 and the

ITS2 regions of P. thornei are very different from those of P. curvicauda.

3.5 Conclusion

After morphological and morphometric analyses, diagnostic ITS and D2-D3 regions of the nematode from four locations were cloned and sequenced, revealing inter and intraspecies variation. The sequencing revealed that the D2-D3 sequences of P. curvicauda collected from these sites confirmed that it was a separate species, one which had been described previously from WA. This conclusion was supported by subsequent phylogenetic analyses of the molecular data. Despite a high level of intraspecific divergence, the ITS sequences also revealed clear separation between P. curvicauda from P. neglectus, P. penetrans and P. thornei. However, there is clearly a need for a more wide spread survey to solve the taxonomic issues of the RLN species present in grainbelt areas of WA.

Chapter 4. Susceptibility of cereals, common crops and pasture species to P. curvicauda

4.1 Introduction

RLNs are the most important group of phytonematodes responsible for causing economic yield losses of winter cereals in Australia. Losses caused by this group of nematodes are increasingly being recognised, particularly under conditions of moisture-and nutrient-stressed soil. Depending on the availability of suitable hosts, moisture stress and soil composition, grain yield losses due to RLNs have been estimated variously from 10% to more than 50% in infested crops and these losses are primarily wheat (Vanstone et al., 1998; Taylor et al.,

2000; Thompson et al., 2008, 2010). RLN infestation can cause significant yield loss with up to 38% in wheat and 18% in barley as revealed from yield loss trial in WA (Collins et al.,

2015). The extent of damage depends on the host plant species and genotype (Wilkinson &

Collins 2014). Moreover, the presence of more than one species in the same field complicates management practices for different RLNs.

Cultivation of preferably resistant or even non-host crops helps in reducing the initial inoculum level and minimise the host damage caused by nematodes. Pratylenchus species have a broad host range making it difficult to find non-host crops for rotation to decrease populations below damaging levels. There is great diversity among the species of

Pratylenchus in terms of pathogenicity (Vanstone et al., 2008). Moreover, among individual crops, the crop cultivars with resistance or tolerance to one species are not necessarily resistance or tolerant to another species of Pratylenchus (Smiley & Nicol 2009). Field pea, lupin and faba bean are poor hosts for P. neglectus but good hosts for P. penetrans, whereas canola is a good host for P. neglectus but not for P. thornei. Oat and barley varieties are also moderately good hosts for P. neglectus (Vanstone et al., 2008). In addition, there are wide

differences in host range and virulence among species of Pratylenchus family. For example, field pea, lupin and faba bean are resistant to P. neglectus, but are susceptible to P.

Penetrans, while oat and barley cultivars are moderately resistant to moderately susceptible to P. neglectus. Conversely, canola is more susceptible to P. neglectus than P. thornei

(Vanstone 2009). Not only that, they might have differences in virulence in regards to climatic variation as well. In Australia, field peas had decreased populations of both P. neglectus and P. thornei (Hollaway et al., 2000; Taylor et al., 2000), while increased populations were found in the U.S.A with same species and host crops (Smiley & Machado

2009). This indicates the type of differences in susceptibility/resistance found for different cultivars of the same crop species grown in different countries.

It is therefore important that responses of important cultivated crop plants to P. curvicauda be assessed to enable effective management. As part of the overall understanding of the biology of this species, it is also important to assess its interaction with wild relatives of cereals in Australia.

4.2 Materials and methods

4.2.1 Sources of plants and germplasm

A total of 61 different genotypes of cereals, pasture plants, canola, legumes and solanaceous plants belonging to five different families were evaluated for their ability to support development and reproduction of P. curvicauda (Table 4.1). The cereals included cultivars and wild germplasms of barley, wheat, oat, sorghum and millet and cultivars of eight different legumes as well as genotypes of the economically important crops soybean, canola, tomato and the Australian native and model plant, Nicotiana benthamiana. Seeds of the wild barley accessions (WA 11724 and WA 11730) were provided by Dr. Chengdao Li

(Western Barley Genetics Alliance, Murdoch University). The wild emmer wheat (TTD 201)

was obtained from Dr. Wujun Ma, Murdoch University. Wild germplasms of cereals (27) obtained from the Australian Grains Genebank (AGG), Horsham, Victoria, Australia were used for this trial. All other seeds including those of the crop and pasture plants, and those of barley cv. Stirling, previously identified as being susceptible to P. quasitereoides (Wilkinson et al., 2014, unpublished data), were obtained from the Nematology Section, DAFWA.

4.2.2 Growing test plants and nematode inoculation

The seeds of all the plant species used were sown directly in pots containing 1 kg of soil consisting of 50% Gingin loam and 50% yellow bricklayers sand. The nematode inoculum suspension was provided by DAFWA, and used to inoculate plants: this inoculum was originally collected from infested soil from glasshouse pot trials. For each plant species, a total of 12 seedlings were inoculated: there were three seedlings per pot with four replicated pots. Fourteen-day-old seedlings were inoculated with 1,000 mixed stage (vermiform) nematodes per pot. Inoculation involved pipetting equal volumes of nematodes suspended in

2 ml of water into 2-3 cm deep holes around the roots of each seedling. The plants were not watered the day before the inoculation, and after that watering were done every other day.

The soil was fertilised with Thrive (Yates) once during the growing period of the experiment at the recommended rate. Susceptibility of the plants to nematode infestation was examined

10 weeks after inoculation.

Susceptibility of the wild cereal accessions was assessed using field soil from

Pingelly, WA (32°32′2.4″S, 117°5′9.6″E). This soil was heavily and predominantly infested with one RLN species (12 nematodes/g of soil), and was used because it provided edaphic factors similar to conditions for possible field infestation.The soil was first sieved using a 1 cm mesh and then mixed homogeneously using a cement mixer.The seeds of test plants were sown directly into the soil and the plants assessed 10 weeks later.The initial nematode

Table 4.1. Crops/cultivars assayed for supporting the reproduction of P. curvicauda

Plant family Common name Cultivar/Accession No. Taxon Plant family Common name Cultivar/Accession Taxon No. Poaceae Wheat Mace Triticum aestivum Poaceae Sorghum Superdan II Sorghum vulgare Wyalkatchem Sweet Jumbo Fortune AGG112151SORG1 Sorghum bicolor Calingiri AGG316510SORG1 Wild emmer wheat TTD201 Triticum turgidum ssp. diccocoides AGG300167WSOR1 Sorghum halepense AGG34043WHEA1 Triticum turgidum diccocoides AGG300263WSOR1 Sorghum Dr.ummondii AGG3697WHEA1 Triticum monococcum boeticum Triticale AGG3959 Triticale sp. AGG15367WHEA1 Millet Nutrifeed Pennisetum glaucum AGG15441WHEA1 AGG108073MILL1 Setaria italic AGG27046 WHEA1 AGG108802MILL1 AGG90350WHEA1 AGG108568MILL1 Panicum miliaceum AGG90383WHEA1 AGG302101MILL1 AGG17458WHEA1 Secale montanum Ryegrass Vortex Lolium multiflorum AGG22454WHEA1 Triticale sp. Fabaceae Field pea Kaspa Pisum sativum AGG22457WHEA1 Triticale sp. Dundale Barley Stirling Hordeum vulgare Chickpea Sona Cicer arietinum Hamelin Howzat WA 11724 Lupin Jennabillup Lupinus albus WA 11730 Narrow Leaf Lupin Mandelup L. angustifolius AGG407588BARL1 Serradella Cadiz Ornithopus sativus AGG400202BARL1 Lucerne Venus Medicago sativa AGG402778BARL1 Hordeum agriocrithon Sub clover Trikkala Trifolium subterraneum AGG407241BARL2 Woogenellup Oat Saia Avena sativa Persian clover Prolific Trifolium resupinatum Dalyup Canola Cobbler Brassica napus Coomallo Brassicaceae Tornado Williams 2332 Forage Brassica - Bannister 2354 Solanaceae Tomato Tiny Tim Solanum lycopersicum AGG701765OATS1 Avena sterilis Tobacco - Nicotiana benthamiana AGG701905OATS1 Avena fatua Soybean - Glycine max Leguminosae Biserrula Casbah Biserrula pelecinus

population in the soil was determined by extracting and taking averages from five 100 g subsamples using the misting apparatus (Tan et al., 2013); the nematodes were collected in

50 ml syringes with silicone rubber connected to the end, sealed with a spring clip.

Nematodes were collected 72 h after the soil was placed on the coffee filters in the mist apparatus, and were counted using a Sedgewick-Rafter counting chamber (Graticules Ltd,

Tonbridge, UK) under a compound microscope. The average initial population was 12 nematodes/g of soil calculated by counting from three replicates of 1mL of nematode suspension from each soil extract. For each test plant, there were three plants per pot with four replicated pots maintained in a glasshouse at 25C, maintained minimal watering to prevent nematodes being washed out of the pots, until nematode populations in the soil and plants were assessed 10 weeks later.

4.2.3. Observation of inoculated plants and nematode infestation analyses

Nematode treated plants and controls were harvested and analysed 10 weeks after inoculation. The pots were not watered for 48 h before plant harvest, to encourage the nematodes to remain in the roots. Nematodes were extracted from roots of the test plants and from infested soil. To extract nematodes from roots, plants were cut at the soil level; the roots were carefully harvested, and the adhering soil was removed by gentle shaking. The roots were then washed lightly with tap water into the pot of soil and thoroughly examined for any visible symptoms of nematode damage. They were then cut into 1 cm pieces and placed on coffee filters in the mist apparatus for nematode extraction (Tan et al., 2013). Nematodes from infested soil were also extracted using the misting apparatus from four replicates of 100 g soil. In both cases, the nematodes were collected from the mister 72 h later and were counted using a 1 ml Sedgewick Rafter counting chambers: from this the total number of nematodes per pot of soil or per root was estimated. After nematode extraction, the roots were oven dried for 48 h at 52°C and weighed. The field soil was predominantly infested 66

with P. curvicauda, the nematode was distinguished from the low numbers of other nematode species using the vulval position of mature females.

The Reproduction factor (Rf) of the nematodes was calculated using the formula Rf=

Pf/Pi, where Pf is the average population of nematodes per pot (i.e. extracted from both root and soil for a cultivar/germplasm) and Pi is the initial population of nematodes used for inoculation or that existed in infested soil before plants were sown (O’Brien 1983). All statistical analyses were performed with SPSS 21.0 and means were separated using Tukey B test at the p<0.05.

4.3 Results

4.3.1 Reproduction of P. curvicauda in test plants inoculated with vermiform stages of nematodes

Because Pratylenchus is a migratory endoparasite which feeds from host roots, but - can leave roots and enter the surrounding soil, the number of nematodes per pot represented those in roots as well as soil at the time of assessment. In general, about 53% of nematodes in both roots and soil were adults, and about 40% were J2 to J4. The occasional eggs extracted from soil or roots of inoculated plants accounted for only 5-7% of the total nematodes counted. When counted, the average number of nematodes in pots of barley cv. Hamelin was two times that of the initial inoculum, indicating that the nematodes developed and reproduced in the plants (Fig. 4.1). Similarly, for six other cultivars of barley (cv. Stirling), sorghum (cv. Superdan II), wheat (cv. Fortune, Wyalkatchem and Mace) and millet, the Rf

(1) indicated there was an increase in the nematode population (Fig. 4.2). Of the 34 different plants tested, 23 sustained nematode populations to varying degrees. Among these, all except the oat cv. Saia (p<0.05) supported significantly fewer nematodes per pot compared to the initial inoculums. The nematode populations in pots of sorghum cv. Sweet Jumbo, oat

cultivars cv.s Williams 2332 and Coomallo and canola cv. Cobbler were up to 25% lower than the initial inoculum. Plants with >50% average reductions in the number of nematodes per pot were mostly non-cereals; they included cultivars of the legumes chickpea, soybean and field pea, and tomato. Pots of one germplasm/wild accession of wheat (TTD 201) and two of barley (WA 11730 and WA 11724) also had fewer nematodes per pot; respectively a

32% and >50% reduction for the wheat and barley accessions (p<0.05) (Fig. 4.1).

Generally, the forage and pasture species did not support growth and reproduction of this nematode species; the average numbers per pot were significantly reduced after 10 weeks, or the nematodes persisted with very low numbers. Except for ryegrass cv. Vortex, for the other species tested, sub clover cv. Trikkala, biserrula cv. Casbah, and the forage brassica there was a reduction in nematode numbers per pot of at least 80% less than the initial inoculum.

No nematodes were present in pots of four species: serradella cv. Cadiz, lucerne cv. Venus and persian clover cv. Woogenellup (Fig. 4.1).

Figure 4.1. The average number of P. curvicauda (Pc) in soil and roots of crop and pasture plants 10 weeks after inoculation with a suspension of 1000 mixed stages of the nematode. Replicates (n=12).

Figure 4.2. Multiplication of P. curvicauda in crops and pasture species in soils inoculated with standard nematode inoculum.

Overall, more nematodes were extracted from roots than from soil; the exceptions were from pots of sorghum cv. (Sweet Jumbo), chickpea (Howzat), canola (Tornado), for which the percentage differences between nematodes from root or soil were not significantly different (p<0.05). For example, although only 10-17% of the initial inoculum remained in pots of the pasture species (e.g. sub clover cv. Trikkala), field pea cv. Dundale, and N. benthamiana, as many as 89-100% were in the roots compared to 45-70% of the total in roots of the more susceptible wheat and barley cultivars. Also for cultivars of the same species with similar numbers of nematodes per pot, the proportions of nematodes in roots and soil varied significantly (p<0.05).

4.3.2 Challenge of wild germplasm of cereals with soil naturally infested with P. curvicauda

Susceptibility of 25 wild accessions of cereals with P. curvicauda was assessed with field soil naturally infested with a high nematode population density. The barley cv.s Stirling and Hamelin, and the wheat cv. Calingiri were included as susceptible controls whereas lupin cv. Jennabillup was used as a non-host control. As an additional control, replicated pots of soil with no plants were used to assess the initial population (e.g. 12,000) so that an Rf could be calculated (O’Brien 1983). After 10 weeks, a quarter of the initial population

(3,046/12,000) remained alive and active in control pots with no plants. Thirteen of the accessions and the wheat cv. Calingiri and barley cv. Hamelin supported nematode growth, development and reproduction of the nematode, because there were significantly more nematodes than were present in the initial infested soil with no plants (p<0.05); these had

Rf3046 between 1.16 and 5.02 (Fig 4.4).

The different wheat germplasm, including cv. Calingiri, generally supported reproduction of P.curvicauda with a Rf, Rf3046 of (1), except for three accessions, namely

AGG34043WHEA1, AGG22457WHEA1 and AGG15441WHEA1. All the barley germplasms assessed were susceptible to this nematode species with Rf1 (Fig 4.4). All the four accessions of millet and three of the four accession of sorghum used for this experiment supported P. curvicauda, except AGG316510SORG1. In contrast, lupin cv. Jennabillup and triticale AGG3959 had the fewest nematodes per pot. The control pots were treated in the same way as the pots with test plants, including watering. The total number of nematodes extracted from roots of all plants was less than that from soil (Fig 4.3). The two genotypes with highest number of nematodes in roots were AGG22454WHEA1 (72%) and sorghum accessions AGG112151SORG1 (88%), whereas, for the rest of the accessions, between 5% and 25% of the total number of nematodes extracted were in the roots, except for the non- host lupin cv. Jennabillup, where all the live nematodes extracted after 10 weeks were in the soil.

Figure 4.3. Numbers of P. curvicauda in roots of cereal germplasms in naturally infested soil (n=12).

Figure 4.4. Multiplication of P. curvicauda in crops and pasture species in naturally infested soil.

4.3.3 Plant response, root structure and nematode development

No brown lesions typical of host responses to RLN infestation were observed on roots of any of the tested plants ten weeks after manual inoculation or planting in the infested field soil. There was a significant variation in the dry root weight of the test plants, ranging from

0.12 to 2.77 g for the inoculated plants and 0.1 to 2.3 g for the germplasms (p<0.05).

Similarly, the level of susceptibility of the test plants to P. curvicauda varied significantly

(p<0.05). The effect of nematode infestation on root weight of individual test plants was not determined directly. However, there was no correlation between root density of the plant genotypes and nematode population in soil or in the roots of these plants at the time of assessment. Notably, the average root weight of the lupin cv. Jennabillup was unusually low.

Seven days after nematode infestation the plants generally had a small tap root and the plants only developed a few lateral roots. In this case, it is possible that the root structure might have affected penetration and feeding of the nematode, resulting in the low number of nematodes retrieved from soil and roots 10 weeks after inoculation. It is also possible that some nematodes feeds ectoparasitically and damaged roots. Perhaps the plant is resistant but intolerant.

4.4 Discussion

Because the life cycle of root lesion nematodes is typically 3-9 weeks in compatible host plants (Castillo &Vovlas 2007), it was expected that the degree of susceptibility of a host to P. curvicauda would be evident by 10 weeks after inoculation. This was indeed the case as compared to the control pots with no plants, the nematode populations increased in some millet, wheat, barley and sorghum cultivars over this period. An increase in nematode populations in wheat, sorghum, oat and barley germplasms or in cultivars indicates that these plants were good hosts of P. curvicauda.

Not surprisingly, the level of nematode reproduction varied with host plant genus and species. It ranged from the solanaceous plants and some cereal lines and cultivars which supported nematode reproduction to legumes and particularly the pastures, sub clover, serradella, persian clover and lucerne, which appeared to be non-hosts. For the latter either no nematodes were extracted from soil or roots of inoculated plants 10 weeks later, or the population was drastically reduced. The multiplication of this nematode species in canola cultivars was in between those of cereals and lupin. Similar reactions have been reported from glasshouse trial for P. neglectus and P. quasitereoides (Collins et al., 2015). In addition, lupin and oat had multiplication factors lower than 1, except for one accession, which was in contrast with multiplication factors for P. quasitereoides greater than 1 on lupin and oat

(Vanstone et al., 2005).

Host plant responses to most RLNs is genotype dependent (Farsi et al., 1995; Jones &

Fosu-Nyarko 2014; Fosu-Nyarko & Jones 2016). The notable variation in nematode multiplication between cultivars of the same crop species or germplasm could reflect different levels of resistance of the plants to Pratylenchus species (Hollaway et al., 2000;

Smiley & Nicol 2009; Wilkinson & Collins 2014; Meagher et al., 2015). Similar differential responses of wheat and oat to other RLNs (e.g. P. neglectus and P. thornei) have been reported in Australia and elsewhere (e.g. North America, Mojtahedi et al., 1992; Hollaway

2002; Ballard et al., 2006). Successful development and multiplication of nematodes in a host plant could also be attributed to inherent plant phenotypes such as root architecture. In this study, the low infestation of lupin may be related to the low root density and apparent lack of lateral roots. Collins et al., (2013) reported similarly low levels of P. quasitereoides infestation of serradella and also lupin, which appears to be resistant to P. penetrans.

For the test plants, the majority of nematodes were extracted from the soil, which showed that the nematodes fed ectoparasitically or that they did enter host roots and would

have fed from host cells, and then exited from the root some time after infection.

Pratylenchus species feed both ectoparasitically and endoparasitically and many motile nematodes can be found in both susceptible roots and the surrounding soil (Zunke 1990).

This feeding behaviour, which appears to create entry and exit wounds in host roots, has been suggested as a primary cause of secondary infection by soil bacteria and fungi, which in turn contribute to the development of browning and lesions typically associated with RLN infestation. However, in this study, there was no browning or lesions in roots of any of the test plants which had been inoculated, or of those planted in the heavily infested field soil after 10 weeks. Although the soil used for direct inoculation had been pasteurised, the field soil was not. If, in fact, infestation of roots with this nematode species does not cause obvious brown lesions on host roots, identification of its presence, damage and associated yield and economic losses, will be more difficult to ascertain. This aspect of host response to this pest needs to be studied further.

The development of host plant resistance provides an economically and environmentally-friendly approach to reducing losses caused by such phytonematodes. This may be achieved either through incorporation of resistance genes into susceptible cultivars or by conventional plant breeding approaches, often involving marker-assisted selection. No major genes conferring resistance to RLNs have been found, and so combinations of minor genes (QTLs) are needed to improve host resistance (Zwart et al., 2005; Schmidt et al., 2005;

Williams et al., 2002; Farsi et al., 1995; Linsell 2013). However, no such work has been undertaken for P. curvicauda, and it is unlikely that any cross-protection will be available using resistance genes identified for other RLN species. For other crops such as tree species, where applicable, resistant rootstocks or break crops can reduce population levels (Jones &

Fosu-Nyarko 2014). In most cases, resistance genes have been sought from wild relatives of cultivated species. For example, this is also the case in that some genes providing resistance

to P. neglectus have been identified from triticale (Farsi et al., 1995). Due to the inherent variability of nematode populations, a number of replicated experiments both in glasshouse and field conditions are necessary to define varietal resistance to a particular species of

Pratylenchus.

4.5 Conclusion

The identification of wild germplasm and crop germplasm with reduced replication of

P. curvicauda would increase understanding of mechanisms of resistance to this nematode, and could provide useful sources of germplasms for breeding resistant lines. The use of biotechnology-based resistance is also an option, with some advances in this area for RLNs.

Knowledge of non-hosts also provides a list of plants that could be used as break crops between growing seasons of susceptible wheat cultivars to reduce inoculum levels and increase yields.

Chapter 5. Attraction, penetration and biology of P. curvicauda

5.1 Introduction

As one of the three most economically important groups of PPNs (Jones et al., 2013), RLNs are mostly polyphagous in nature. Feeding is usually restricted to the root cortex (Castillo &

Vovlas 2007). Interactions of Pratylenchus spp. with roots involves attraction, penetration, feeding and migration in roots and also leaving and re-entering roots (Linsell et al., 2014).

RLNs use their mouth stylet to puncture root epidermal cells and may feed ectoparasitically, but then enter roots and become endoparasites. Attraction of both juvenile and adult RLNs to roots is thought to be influenced by gradients of compounds present in the rhizosphere or secreted root cells or the root cap (Fosu-Nyarko & Jones 2016). Information on differences in biology and penetration of Pratylenchus in resistant and susceptible hosts is limited.

Although the nematodes P. neglectus and P. thornei have been reported to penetrate resistant and susceptible hosts equally (Talavera & Vanstone 2001; Farsi et al., 1995), resistance responses then appear to restrict motility and reproduction (Baldridge et al., 1998; Prasad et al., 1999; Rich et al., 1977). Whether a plant is resistant or susceptible depends both on hosts and on the nematode species. For instance, the cultivars of wheat which are resistant or tolerant to P. thornei do not necessarily express resistance to P. neglectus, which indicates that responses to each species are genetically independent (Smiley & Nicol 2009).

Resistance of a specific crop/cultivar to nematode infestation is dependent more on its genotypes than on the number of nematodes that penetrate roots initially (Linsell et al.,

2014). Because of this genotype dependent response, information on interactions of economically important species of Pratylenchus (e.g. P. neglectus, P. thornei, P. quasitereoides or any new species) needs to be determined. In particular, whether a particular genotype is susceptible, resistant, tolerant or intolerant.

RLNs have five stages in their life cycle including eggs, all the motile stages are infective. Thus J2, J3, J4 and adult stages are parasitic and both penetrate and leave host roots. In favourable condition RLNs can complete 3-6 generations per growing season

(Taylor et al., 2000), depending on species and environmental conditions these complete the life cycles in 45-65 d, (Smiley & Nicol 2009). Study of life cycles of Pratylenchus species have mainly been done under controlled conditions, with life cycles at higher temperatures and in susceptible hosts. For instance, the life cycle of P. penetrans can be completed in 22 d at 30°C in red clover, while its life cycle was twice as long (46 d) at 17°C (Mizukubo &

Adachi 1997). For P. vulnus, one generation was completed in 26 d on carrot discs at 26°C

(Chitimbar & Raski 1985).

This chapter describes (i) the life cycle of P. curvicauda in roots of a susceptible wheat cultivar in soil, and (ii) early interactions of this nematode species with some resistant and susceptible cereals and pasture species. The data generated can contribute to understanding their biology and to improve their control.

5.2 Materials and methods

5.2.1 Nematode inoculum

The nematodes used in the work in this chapter were extracted from soil collected from Pingelly and nematodes were extracted using the misting apparatus and were stored at

4°C. Nematodes used for the life cycle study were J2 and J3 stages. These were separated from other stages using a series of sieves: 38, 30 and 20 µm diameter mesh. The adults were separated from juvenile mixed stages using 38 µm sieves, whereas passage through the 30

µm and 20 µm sieves was used to separate J2 and J3 stages. The body lengths of 12 nematodes of each stage were measured. For this, nematodes were hand picked using the fine tipped feather, and placed in a drop of water. To measure body length, nematodes were

placed on a slide, and were gently heated to immobilise them. The microscopic image scale was used to measure the length. The body length of J2s was 240-300 µm, with an average of

267 µm, the J3s were 300-372 µm long with an average of 327 µm and the adults were 528-

584 µm long, averaging 560 µm. In the sample collected as adult nematodes, some looked mature, but without a vulva indicating that there might still be same J4 stages of the nematode in this fraction.

5.2.2 Attraction of P curvicauda to host

The same cereal genotypes and pasture species used for the penetration assay were also used to study attraction of nematodes to roots. Seven-day- old seedlings were transferred to Petri dishes with lines drawn underneath to measure movement of nematodes. For each plate, a seedling was placed in the middle with a thin layer of 0.4% water agar, and nematodes (600/root) were placed 10 mm from the root (Fig 5.1). The number of nematodes which had migrated towards the roots was counted 2, 4, 6 and 12 hours after inoculation

(hai), and expressed as a percentage of the initial number of nematodes added. There were four replicates for each crop genotype.

5.2.3 Penetration of P curvicauda to host

Fifteen different genotypes of cereal and pasture species determined as resistant and susceptible to P. curvicauda were used in this study. The germplasm was selected based on their suitability as hosts of this nematode as determined in Chapter 4. Seeds of all cultivars were pre germinated on moist filter paper at room temperature for 3 d and then transferred to pots containing 100 g of pasteurised sand. The nematode inocula were added just on the roots

@450/plant, and covered gently with sand. Plants were watered sparingly to maintain soil moisture, but without washing nematodes away. Roots were harvested at 12, 24 and 48 h, and

7 and 14 days after inoculation (dai) to determine the number of nematodes that had

penetrated the roots. Each root infection assay was replicated four times per

cultivar/germplasm.

Figure 5.1. Attraction of P. curvicauda to host roots in a Petri dish (R=Root, N=Nematode).

5.2.4 Life cycle of P curvicauda

Wheat cv. Calingiri, a susceptible host for this nematode species, was selected for this

study. The seedlings were initially grown in corrugated trays with vertical columns (Fig 5.2).

Three days after seed emergence in the trays, seedlings were inoculated with J2 and J3 stages

of the nematode @250/plant. Each root was covered with a small amount of fine pasteurised

sand after inoculation and plants were carefully watered to keep the sand moist. After 7 d,

inoculated seedlings were transferred to trays (Fig 5.2) and the trays were transferred to a

glasshouse to facilitate plant growth up to the end of the experiment. The temperature of the

glasshouse was 23°C. The four replicates of roots were harvested and the stage development

was determined at 5 d intervals from the 15 to 60 dai.

Figure 5.2. Seedlings in corrugated trays (left and middle) and transferred seedlings in tray pot (right).

5.2.5 Staining and counting of nematodes

In order to follow penetration, migration and reproduction of P. curvicauda in

inoculated plants, roots were washed gently after harvesting and stained with acid fuchsin

(Byrd et al., 1983). The roots were soaked in commercial bleach (NaOCl) solution diluted to

1.5% for 5 min followed by washing with running water to remove residual NaOCl. The

roots were heated to boiling in a beaker containing of 1 ml stain (3.5 g acid fuchsin, 250 ml

acetic acid, and 750 ml distilled water) in 30 ml of water for about 1 min. The roots were

cooled to room temperature and washed in running water to remove excess dye. They were

then mounted on a slide to determine the number of nematodes inside the roots using a

compound microscope (Olympus BX51).

5.3 Results

5.3.2 Attraction of nematodes to resistant and susceptible roots

The attraction assay was undertaken using the same resistant and susceptible crop

genotypes as in the penetration assay. The attraction of nematodes towards roots was

calculated as a percentage of the initial number of nematodes used. The number of nematodes

attracted to roots of all cultivars/genotypes increased with time, as was observed for the penetration assays (Fig 5.3)

The highest percentage (~40%) of nematodes were attracted to susceptible wheat genotype (AGG17458WHEA1) and also to three resistant genotypes of triticale (triticale

AGG3959), wheat (AGG15441WHEA1) and millet (AGG108568MILL1) (Fig 5.4) and they were statistically similar (P=0.05). Conversely, the lowest percentage (7%) of nematodes attracted to cereal plants was to oat (AGG701905OATS1) roots, which was in fact a suitable host with Rf=2 for this nematode species. Overall, 7-13% of the nematodes present were attracted to the roots of the pasture species after 12 h.

Figure 5.3. Attraction of nematodes to roots: a. movement of nematodes towards root hair zone and upper part of roots after 6 h; b. nematodes around a root after 6 h (wheat cv. Calingiri); c. nematodes around a root after 2 h (AGG17458WHEA1).

Figure 5.4. Nematodes attracted towards roots of cereal and pasture plants after 2, 4, 6 and 12 h. The designations are: S-susceptible, R-resistant, HS-highly susceptible, HR- highly resistant (refer to Chapter 4). For each time period, bars with the same letter did not differ significantly (P=0.05) according the Student's t-test.

5.3.1 Penetration of nematodes in resistant and susceptible genotypes

Nematodes were not present in roots at 12 and 24 h after inoculation. Some nematodes were present in roots of wheat (cv. Calingiri, AGG17458WHEA1,

AGG15441WHEA1), sorghum (AGG316510SORG1), oat (AGG701905OATS1) and triticale (triticale AGG3959) 48 h after inoculation, but not for lupin (Jennabillup), wheat

(AGG27046WHEA1), sorghum (AGG300263WSOR1), millet (AGG108568MILL1) or any of the pasture species. The greatest number of nematodes at each time point was recorded from the triticale (AGG3959) roots and was significantly higher (P=0.05) than other cultivars/genotypes used (Fig 5.5). This is of notable since the results of the host range study indicated that triticale was a poor host with a reproductive factor (Rf) of only 0.26. Two

Figure 5.5. Nematodes inside roots of wheat (AGG17458WHEA1) (left) and triticale (AGG3959) (right) after 14 dai, staining with acid fuchsin.

susceptible wheat genotypes, Calingiri (used as susceptible control) and AGG17458WHEA1

(Fig 5.6) had the second highest number of nematodes in roots, followed by one resistant

genotype of wheat (AGG15441WHEA1) and one resistant sorghum germplasm

(AGG316510SORG1), although there were not significant (P=0.05) differences between

them.

Conversely, the lowest percentage of nematodes was in roots of two susceptible oat

(AGG701905OAT1) and wheat (AGG27046WHEA1) genotypes 48 h to 14 dai. Although

few nematodes entered roots of some pasture species, such as ryegrass cv. Vortex and sub

clover cv. Trikkala, no nematodes were found in roots of the other pasture species tested (sub

clover, persian clover, biserrula) or in lupin (cv. Jennabillup), which was used as resistant

control. Overall, a maximum of approximately 20% of the inoculated nematodes penetrated

roots of triticale (HR-highly resistant) at 7 dai, although the number decreased at 14 d (Fig

5.6). A similar result was found for the resistant wheat genotype (AGG15441WHEA1), and

for sorghum (AGG316515SORG1) genotype. In contrast the number of nematodes increased

in roots of susceptible wheat (Calingiri), sorghum (AGG300263 WSORG1) and the highly

Figure 5.6. Percentages of P. curvicauda in roots of cereal and pasture species 48 h to 14 dai. The designations are: S-susceptible, R-resistant, HS-highly susceptible, HR-highly resistant (refer to

Chapter 4). For each time period, bars with the same letter did not differ significantly (P=0.05) according to the Student's t-test.

susceptible wheat (AGG17458WHEA1) at 14 d compared to 7 d, which indicated the attractiveness of nematodes of these host roots.

5.3.3 Life cycle of P. curvicauda in wheat

A total of 30 individual nematodes of each stage (J2, J3, J4 and adult) were measured from each harvest to record stage development in roots. An increase in average body length was then measured after 25 d. Fourth stage juveniles were present on 30 d, and adults on 35 d. Eggs were found at 40 dai (Fig 5.7). After this period the average body length of the nematodes present dropped as more juveniles hatched from the eggs (Fig 5.8 & 5.9).

Figure 5.7. Days required for J2-J3 of P. curvicauda to complete its life cycle in a susceptible wheat cv. Calingiri (not to scale).

Figure 5.8. The average body length of the nematode P. curvicauda in wheat cv.

Calingiri roots at different days after inoculation.

Figure 5.9. Life stages of P. curvicauda (J2-J2) in wheat roots stained with acid fuchsin from 15-

50 dai.

As RLNs are generally characterised as inducing lesions in infected roots resulting from migration and feeding, the roots were checked thoroughly at each harvest to assess if

any lesion was present. However, typical lesions were not observed in any of the roots.

However, there were dark brown patches in a few roots 55 d after infection, but these were not typical symptoms of RLNs. Staining and closer examination by microscopy indicated the presence of a fungus in some roots (Fig 5.10).

Figure 5.10. Brown discoloured wheat roots at 55 dai; b-d: Fungal structures found in roots of wheat cv. Calingiri.

5.4 Discussion

This research shows clearly that early attraction and penetration of P. curvicauda was not affected initially whether or not the host was resistant or susceptible, unless it was a non- host. Both the rate of movement of nematodes towards roots and the penetration increased 90

with time after inoculation. RLNs probably use a combination of their mouth stylet and gland secretions to enter and migrate in host root tissues. Since RLNs are polyphagous, they are likely to possess a redundant variety of cell wall modifying enzymes enabling them to migrate in host roots of many species (Fosu-Nyarko & Jones 2016). Hence, most of the nematodes were able to overcome any initial physical barriers of both resistant and susceptible crops/genotypes. In this study, it was found that nematodes had no preferential site for penetration and most nematodes were found in more mature regions of roots, not particularly at the root tip zone. Similar penetration patterns were evident for P. thornei in chickpea roots (Castillo et al., 1998), whereas this contrasted with P. penetrans which was attracted to penetrate in the root elongation zone (Zunke 1990). However, P. penetrans also penetrated through root hairs when inoculated to strawberry roots (Kurppa & Vrain 1985).

It is interesting that the P. curvicauda was attracted to and penetrated roots of susceptible and resistant cultivars of host plant species equally. This suggests the resistance must be manifested by mechanisms other than any operating during the initial interactions. In fact, similar results have been found for P. neglectus and P. thornei penetrating wheat and chickpea cultivars/genotypes (Linsell et al., 2014; Farsi 1996). P. zeae similarly invaded both resistant and susceptible sugarcane clones 12 and 24 h after inoculation (Khatiresan & Mehta

2002b). This result did not apply for non-host species, which were not attractive to the nematodes.

The root development of different seedling genotypes varied. For example, the root systems of triticale AGG3959 and wheat cv. Calingiri grew faster than that of WHEA1 AGG

17458, so more roots might attract more nematodes. Conversely, lupin had very poor root development with only a tap root present at the same period. Similar poorly developed roots were found for the pasture plants ryegrass, sub clover, biserrula and persian clover, which were not suitable hosts for this nematode species. In those cases, where fewer nematodes

were present in roots after 14 d compared to 7 d, this reflects the migratory nature of

Pratylenchus, and since the penetration assays were conducted in soil, nematodes could exit roots and return to the soil.

RLNs generally complete their life cycle within 3-9 weeks, influenced by host and nematode species. However, the developmental rate also depends on environmental factors such as temperature, moisture and host (Taylor et al., 2000). This study provides the first information on the length of the life cycle of P. curvicauda which can complete one generation by 35-42 d at 23°C in wheat cv. Calingiri. Comparable data for P.thornei in wheat also depends on temperature, which reproduced at 20-25°C on all wheat cultivars, but it did not reproduce at 30°C at all and a very little at 15°C (Thompson et al., 2015). Results for the test nematode showed that the greatest number of nematodes were present at 40 d, after which numbers dropped presumably as nematode left the roots. Similar observations have been made for P. scribneri where the older stage (J4 or adult) were more likely to leave roots than J2 and J3 (MacGuidwin 1989).

In soils, RLNs are capable of feeding as ectoparasites and able to lay eggs in roots or outside roots in soil. In in vitro studies, although most vermiform stages and eggs were in roots of tomato, soybean and corn, many individuals of P. agilis appeared to be able to feed and complete their life cycles ectoparasitically, by feeding only on root epidermal cells and root hairs (Rebois & Huettel 1986). Remarkably for P. penetrans, eggs have been observed as early as 5 dai in strawberry (Townshend 1963b), but this result may have been obtained because the nematode inoculum was not staged accurately.

The body length and other morphometric values in fixed specimens were usually smaller than those of live specimens as found in P. hippeastri (Inserra et al., 2007). In this case, boiling the roots during staining might cause shrinkage in body length of nematodes in roots. For P. penetrans, J2s were reported as 225 µm, J3 306 µm, J4 was 408 µm and adult 92

417-477 µm long, respectively. The length of various stages also could overlap depending on the state of moulting, as found in P. penetrans (Townshend 1991).

5.5 Conclusion

The work in this chapter focused on the biology of P. curvicauda, particularly on how it responds to resistant or susceptible hosts in terms of attraction towards roots and penetration. Several cultivars/germplasms of cereal and pasture species were examined in vitro for their susceptibility to root penetration. Although nematodes were not attracted to non-hosts, the effectiveness and penetration of poor or good hosts was similar. This indicates that susceptibility or resistance is based on other host plant properties. The life cycle of this species in wheat at 23°C was determined at 35-42 d.

Although these studies were done on cultivated species, it would be useful to know whether native species are also hosts that could provide a reservoir for new infestations, and whether this species is endemic or has been introduced to WA.

Chapter 6. Histological responses of susceptible and resistant hosts infested with P. curvicauda

6.1 Introduction

As migratory endoparasites which do not induce defined feeding sites, histological changes induced in roots on infection with RLNs have been less studied than those caused by sedentary endoparasitic nematodes. After attraction to the root surface, Pratylenchus spp. infection process involves penetration of the epidermis, feeding from individual cells and migration within root tissues (Linsell et al., 2014). Attraction of RLNs, both juveniles and adults, to roots is thought to be a response to gradients of compounds present in the rhizosphere, mainly originating from plant roots (Fosu-Nyarko & Jones 2016). Migration through root cells causes cellular damage in root tissues which subsequently affects uptake or transport of nutrients and water through roots. This migration also provides an avenue for entry by other soil microbes (soil fungi, bacteria) resulting in secondary infections (Castillo &

Vovlas 2007).

RLNs use their mouth stylet to puncture root epidermal cells. The importance of the products of parasitism genes expressed in the oesophageal gland cells and secreted through the stylet into host cells and tissues represents an evolutionary advance and adaptation for plant parasitism by nematodes (Davis et al., 2000). The stylet secretions play a vital role in nematode penetration and migration through root tissues. Depending on the genus, the secretions are also vital for the modification and maintenance of root cells as feeding cells, the formation of feeding tubes, and/or digestion of host cell cytoplasm to facilitate nutrient acquisition by the nematode (Hussey 1989).

PPNs cause important changes in the hosts they infect. These changes start at the cellular level. The initial interaction between the migratory and sedentary endoparasites and the host may be similar; however, as the interaction develops, there are significant differences 94

in host responses to different nematode species, which ultimately affect symptoms of infection, plant growth and development. The changes at the cellular and or at structural level, in root tissues invaded by RLNs have only been studied for a few species, and mostly for P. penetrans (Townshend et al., 1989; Vieira et al., 2107). Since, alterations in infested cells have not been studied in any cereal roots, the aim of this study was to examine the histological and ultrastructural responses induced by entry, migration and feeding of P. curvicauda in wheat and barley roots.

6.2 Materials and methods

6.2.1 Preparation of nematode inoculum

The nematode inoculum used in this study was extracted from Pingelly soil and the nematodes were extracted using the mist apparatus.

6.2.2 Growing of plants and inoculation of nematodes

The wheat cv. Calingiri and barley cv. Hamelin were selected as susceptible hosts and a wild species of barley was selected as a resistant host to undertake this study. Seedlings of each cultivar were grown in corrugated trays as described in Chapter 5 (Fig 5.2). Three day old seedlings of each cultivar were inoculated with nematodes suspended in water, 150-160 nematodes per root and the roots were covered with pasteurised fine sand. Four replicates were established for each cultivar. The nematodes were allowed to penetrate and feed in the roots for up to 15 d. Control plants not inoculated with nematodes were grown similarly for the same period. Roots were collected at different times after infection and washed thoroughly to remove any adhering particles of sand.

6.2.3 Staining of nematodes

To follow migration of nematodes in roots at different time points, the roots were stained with acid fuchsin (Byrd 1983) following the protocol described in Chapter 5. 95

6.2.4 Harvesting and fixation of roots for light microscopy (LM) and transmission electron microscopy (TEM)

The nematode inoculated roots were fixed for microscopy 3, 7 and 15 d after inoculation. The roots were cut in 3% glutaraldehyde in 25 mM sodium phosphate buffer (pH

7.0) and fixed overnight, then removed by washing with phosphate buffer (0.025M) (pH 7).

In initial experiments, paraffin embedded sections were prepared to locate the nematodes in roots. The roots pieces were embedded in paraffin wax. For LM, sections were cut 10 µm thick using rotary microtome (LEICA RM2235) in the form of a ribbon. The ribbons were cut and mounted on slides, which were kept in an incubator at 40°C for 24 h. The sections were stained with a mixture of 1% Azur II, 1% methylene blue in 1% sodium tetraborate

(Richardson 1960). LM sections were examined and photographed with a compound microscope (Olympus BX51).

Processing of roots for TEM involved fixing roots with 3% glutaraldehyde in 25 mM sodium phosphate buffer (pH 7.0) overnight, and washing with 0.025M phosphate buffer (pH

7.0). Since RLNs do not cause obvious symptoms of their presence, the inoculated roots were post fixed with (OsO4) (1%) in 0.025 M phosphate buffer (pH 7.0), for 2 h at room temperature in a fume cupboard. The OsO4 was replaced with 0.025M phosphate buffer (pH

7.0) by two changes at 15 min intervals. Osmium fixation stained nematodes black and so helped to locate nematodes in roots under a dissecting microscope. Selected parts of root which contained nematodes were then embedded in Spurr’s resin following a standard protocol. The glutaraldehyde/osmium fixed root pieces were dehydrated in a graded series of acetone (15 min each (v/v) 30%, 50%, 70%, 90% and 100%) and infiltrated with Spurr’s resin. The specimens were embedded in fresh Spurr’s resin at 60°C for 24 h. Approximately

90 nm thick sections were cut on a microtome (Reichert - Jung Supercut Programmable

Microtome 2050) using a glass knife and captured on 200 mesh copper grids. Sections on

copper grids were stained with 4% uranyl acetate for 10 min and 3% lead citrate for 5 min

and examined at 100 kv using a TECNAI 12 transmission electron microscope.

6.3 Results

To ensure that the nematodes had infected the roots under the conditions used, the

migration of nematodes at different time points was followed by acid fuchsin staining (Fig

6.1). Nematodes were not found in the roots in the first 24 h. At 3 d and longer, the number of

nematodes in roots increased, and was distributed predominantly in cortical cells (Fig 6.1).

Figure 6.1. P. curvicauda in wheat cv. Calingiri roots at different times after inoculation: a. 3 dai; b. 7 dai; c. 15 dai.

Light micrographs of both longitudinal and cross sections of root tissues revealed that

P. curvicauda were present inside wheat and barley roots usually in cells of the cortex (Fig

6.1 & 6.2). Nematodes were usually aligned parallel to the stele as they penetrated transverse

walls of successive cells in the root cortex. The surrounding cells sometimes had thickened

walls (Fig 6.2 & 6.3).

Fig. 6.2. Nematodes lying on cortical cells in: a. Longitudinal paraffin section of wheat cv. Calingiri roots at 3 dai; b. wheat cv. Calingiri roots at 7 dai; c. barley cv. Hamelin roots at 7 dai; d. wild barley roots at 7 dai. (N=nematode).

Fig. 6.3. Light micrographs of nematodes in the cortex of wheat roots at 3 dai : a. P. curvicauda in cortical parenchyma (p=pericycle, e=endodermis, nu=nucleus, N=nematode), deformed endodermis, thickened cell walls in surrounding cells (cross section); b. longitudinal section of roots showing collapse of cytoplasm (marked with arrow), and enlarged nuclei (longitudinal section).

The light microscopy of infested roots clearly illustrated intra-cellular migration of nematodes (Fig 6.4). The cellular responses and feeding behaviour of this nematode appeared to be much the same for both resistant and susceptible roots of wheat or barley. Nuclei of cells of root close to nematodes appeared enlarged (hypertrophied) and there were large cavities in adjacent cells when nematodes had moved from one cell to another (Fig 6.4).

Figure 6.4. a & b. Paraffin sections of wheat cv. Calingiri roots at 3 dai showing intra- cellular migration of P. curvicauda (nu=nucleus; N=nematode); c. Control roots without nematode (longitudinal sections).

In order to identify which fixed root segments contained nematodes, for TEM studies, double staining with glutaraldehyde and osmium (OsO4) showed where the nematodes were located in the roots (Fig 6.5).

Fig.6.5. Nematodes in glutaraldehyde/OsO fixed roots under a dissecting microscope. a. 4 wheat cv. Calingiri 3 dai; b. wheat cv. Calingiri at 7 dai; c. Barley cv. Hamelin at 7 dai.

In TEM sections of infested roots, cell and tissue damage was evident in all wheat and barley inoculated with this Pratylenchus species. Affected cortical cells in infested roots were devoid of cytoplasmic contents and had distorted cell walls (Fig 6.6 c). Large cavities were visible in infested root cells and the cytoplasm of affected cell was degraded (Fig 6.6 d).

The presence of this nematode in root tissues not only affected invaded cells, but similar effects were also observed in cells adjacent to those directly damaged by nematodes, with pronounced alterations in cell morphology and content, including accumulation of granular material in the cytoplasm (Fig 6.6 d). In some sections, the cortical cells appeared to be distorted or compressed which could be a result of loss of turgidity in cells and their collapse after nematode feeding and migration (Fig 6.6 c).

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Figure 6.6. Light and electron micrographs of infested wheat (cv. Calingiri) roots at 3 dai. a. LM of infested roots; b. TEM showing cells containing nematodes (N), but otherwise cells devoid of cytoplasmic contents; c. deformed and compressed phenotype of adjacent cells; d. granular material in the cytoplasm, cells with intact vacuoles (V) and tannin (t) deposition in infested root cells.

Cortical cells which contained nematodes usually lacked cytoplasmic contents, as shown in inoculated wheat roots 7 d after nematode inoculation (Fig 6.7 a & b). In addition, the same images show the cytoplasm in adjacent cells lacked integrity, the vacuolar membranes were contracted or distorted and most organelles and membranes were severely damaged. Other cellular level changes in root cells included condensation of chromatin in nuclei and compression or distortion of endodermal cells (Fig 6.7 c).

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Figure 6.7. TEM of wheat (cv. Calingiri) roots at 7 dai infested with P. curvicauda . a & b. adjacent cell devoid of cytoplasmic content, deformed plasma membrane (p= pericycle, e= endodermis, N= nematode); c. deformed endodermis, nucleus with condensed chromatin; d. cross section of a control root without nematode infection

The effect of P. curvicauda in barley cv. Hamelin and wild barley showed similar alterations in cellular structures as found in infested wheat roots. The nematodes were also present in cortical parenchyma cells (Fig 6.8 a & d). The infested cells exhibited disorganised cytoplasm and cytoplasmic contents, in some sections adjacent cells were empty or present organelles appeared damaged (Fig 6.8 b; Fig 6.9 a & b). Some cells had ruptured cell walls indicating that nematodes had migrated through those cells (Fig 6.8 c).

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Figure 6.8. Barley (cv. Hamelin) roots at 7 dai infested with P. curvicauda. a. light micrograph of root; b. nematode inside cell devoid of cytoplasmic contents with disorganised cytoplasm in adjacent cell; c. ruptured epidermal cell wall (as indicated by arrow) (N=nematode).

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Figure 6.9. Wild barley roots at 7 dai infested with P. curvicauda. a & b. compressed cells with degeneration of cell contents; c. nematode showing vulva; d. control cells without nematode infestation.

6.4 Discussion

Plant cells parasitised by different nematode species are modified in diverse ways.

Cellular changes due to nematode parasitism have been categorised as either destructive or adaptive cell modifications (Dropkin 1969). Adaptive cell changes include those in which longer term feeding sites are induced, such as giant cells induced by RKNs and syncytia by

CNs. The formation and function of giant cells and syncytia have been studied in

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considerable detail (Jones & Northcote 1972a; Jones & Payne 1978; Jones 1981; Jones &

Goto 2011; de Souza et al., 2017). In contrast, for nematodes which cause destructive cellular changes there is much less information. Destructive changes range from removal of the contents of individual cells by feeding nematodes to more extensive destruction of cells and tissues. This chapter provides the first ultrastructural study of P. curvicauda in wheat and barley roots.

Acid fuchsin staining provided an overview of the numbers of nematodes present, and their location in the roots. Since there were no nematodes in roots before 24 h, before this time nematode could be locating roots or feeding ectoparasitically. As for other RLNs, this nematode often curled over in a cell, and in heavy infestations there could be several nematodes in the same cell.

In terms of cellular changes, the response of wheat and barley to this Pratylenchus species was similar to those described for P. penetrans in strawberry, alfalfa and lily (Vieira et al., 2017; Townshend et al., 1989; Zunke 1990a; Acosta & Malek 1981). The nematode infested wheat and barley roots showed alterations in cells including: disintegration of cytoplasm, malformed cells, compression of cells next to penetrated ones and also the appearance of hypertrophied nuclei in some cells. Cells penetrated by the nematode contained only cytoplasmic debris which was composed largely of degenerated organelles and condensed cytoplasm (Townshend et al., 1989). The degradation in epidermal and cortical cells was the result of both of migration of Pratylenchus within cells of infested roots and feeding damage. Similar damage and changes have been reported in other host-Pratylenchus interactions (Vieira et al., 2017). There was usually a large cavity formed in infested root cells, and this was visible in sections examined both by LM and TEM.

Both LM and TEM showed P. curvicauda did not appear to invade the endodermis and vascular tissues. This differs from CNs or RKNs which form syncytia or giant cells 105

associated with vascular tissues of infested roots (Rodiuc et al., 2014). The endodermal cells looked shrunken and distorted, which could possibly have protected or prevented invasion of the vascular bundle by the nematodes. Limitations of RLNs to cortical tissues may protect the host plant from more major damage, and enable water and nutrients to continue to be transported to the aerial plant tissues. Electron micrographs showed tannin deposition in infested root cells, which is usually associated with cytoplasmic disorganisation and lack of membrane integrity (Vieira et al., 2017; Townshend et al., 1989). In addition, apparently thickened cell walls, observed in LM, could possibly reflect deposition of condensed and granular cytoplasm against the walls. Tannin deposition is often observed as a defence response stimulated by plants against pathogen attack (Lattanzio et al., 2006).

Although penetration, migration and subsequent development of the nematode in wheat and barley cultivars/genotypes reported as resistant, tolerant, intolerant or susceptible does not reveal differences in infested roots. Cereal cultivars do have the capability to tolerate some level of damage from nematode infestation. In this respect, it may be that a difference between susceptible and tolerant cultivars is that damage is limited to cortical cells in tolerant cultivars, whereas perhaps vascular cell damage in intolerant cultivars reduces yields more by interfering much more with vascular delivery of water and nutrients to the top of the plant.

Also root vigour and architecture might lead to more roots escaping nematode infestation in tolerant cultivars.

As for other PPNs, RLNs possess putative secreted proteins which act as effector molecules which are needed for successful parasitism (Haegeman et al., 2011; Nicol et al.,

2011; Fosu-Nyarko et al., 2015). The resistant and susceptible genotypes of wheat and barley used were determined based on reproduction of P. curvicauda. However, the genotypes studied all exhibited similar responses to this particular species of Pratylenchus at different time points after inoculation. Although the molecular basis of Pratylenchus-host interaction

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mechanisms has still to be studied in detail, gene expression is thought to be changed more in resistant cultivars than susceptible cultivars of the same crop (Backiyarani et al., 2014; Yu et al., 2015). Invasion by Pratylenchus spp. triggers different signalling pathways during penetration and establishment of hosts (Fosu-Nyarko & Jones 2016; Vieira et al., 2017).

Infestation by P. coffeae showed expression of defence genes and products of the phenylpropanoid pathway which lead to production of secondary defence compounds

(Backiyarani et al., 2014; Yu et al., 2015).

6.5 Conclusion

In this chapter, LM and TEM studies of host tissue of infested wheat and barley roots were studied after inoculation with P. curvicauda. The nematodes were only found in epidermal and cortical cells. The main ultrastructural alterations were visible in cortical cells, and to the endodermis. Cells penetrated by nematodes, and cells next to them, had damaged or modified cytoplasm. Cell organelles were disorganised and cells distorted. Hypertrophied nuclei, sometimes with dense chromatin were present together with degeneration of mitochondria and endoplasmic reticulum. Both resistant and susceptible host reactions appeared similar.

A more detailed microscopic study employing techniques such as differential staining to identify specific components (e.g. starch, lignin etc.) or using advanced techniques such as

X-ray analyses to look for changes in inorganic ions, or scaning electron microscopy of cut roots, which would provide a 3-D view of infected tissues, would further enhance understanding of this host-pathogen interaction.

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Chapter 7. General discussion

At the start of this project, the principal aim was to develop methods to identify and study the biology and host interactions of a root lesion nematode species, recently described as P. quasitereoides (Hodda et al., 2014). It was also thought that this nematode was a new RLN species endemic to Australia and that it is an economically important pest of cereal crops grown in the state. The original nematode inocula provided for this project (by DAFWA) were described as P. quasitereoides, but as the project progressed, it became clear that the taxonomic description of P. quasitereoides did not fit the features of the nematode which was being studied. As a result of the work described here, it now appears that this RLN is in fact

P. curvicauda. Thus, at the start of the project, the specific objectives of the research were to fully characterise P. quasitereoides in terms of taxonomic, molecular and biological aspects.

However, following the morphometric measurements, morphological, and molecular analyses, the species was found to be P. curvicauda, which had been described earlier in samples from WA (Siddiqi et al., 1991). Hence, the objectives of the research were reframed as follows.

1) To undertake morphological characterisation of P. curvicauda

2) To study the genetic relationship of P. curvicauda with other Pratylenchus spp. present in

Australia (P. neglectus, P. thornei P. penetrans, P. quasitereoides).

3) To study its life cycle, biology, attraction to and penetration of resistant and susceptible hosts

4) To study the host range of this species

5) To understand histological and ultrastructural changes which occurred in susceptible and resistant roots of host plants infected by this species of Pratylenchus.

Overall, the main objectives of this thesis were achieved successfully. The major findings are discussed in the general context below.

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7.1 Morphological features of the undescribed Pratylenchus species from four paddock soils of WA

The initial study of the morphology of the species of Pratylenchus provided for this project did not fit the published data of P. quasitereoides, this prompted the need for more study of this species using morphology and morphometric parameters. Morphometric studies alone can be challenging for identification of Pratylenchus species, because the genus has relatively few defining characters which can be used to separate one species from another, there are also some overlapping morphological features.

Both the morphological features of the nematodes and morphometric measurements suggested that the nematode species from four different locations, and in particular samples from Pingelly, were not P. neglectus, P. penetrans, P. thornei or P. quasitereoides. Further morphometric and morphological analyses conducted by an expert in plant nematode taxonomy, Dr. M. R. Siddiqi, identified the nematodes under study from WA as P. curvicauda.

Therefore, from a practical point of view, it is of paramount importance to solve the taxonomic issues related with Pratylenchus species in WA to avoid misinterpretation of data from screening of new wheat, barley or other crop varieties for responses to a particular species of Pratylenchus.

7.2 Genetic variation of P. curvicauda with other Pratylenchus populations in WA

Initial examination of RLNs from soils or cultures confirmed that P. curvicauda differed in morphology from P. neglectus, P. penetrans, P. thornei and. P. quasitereoides.

Standard diagnostic PCR primers were used to amplify ITS regions of about 900 bp for all the four species. Since the ITS sequence of P. quasitereoides had not been studied by Hodda

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et al. (2104), rather they had amplified the D2-D3 expansion region, this region was also amplified and it generated bands of ~300 bp for for all the four species in this project.

When the amplicons generated were sequenced, for the ITS regions, cloned sequences from P. penetrans (930-933bp) were most similar in length to those of P. curvicauda (906-

933 bp), with amplicons of the ITS regions from P. neglectus (825-826 bp) and P. thornei

(826 bp) being substantially shorter. There was also more variation in length of amplicons of

P. curvicauda from four different locations in WA, indicating perhaps that geographical separation had resulted in population divergence, and that P. curvicauda had in fact been present in WA soils for a long time period, and that it could be another endemic species along with P. quasitereoides.

When ITS sequences of the four RLNs studied were compared with those from other locations, including the use of phylogenetic relationships, there were substantial differences in the respective sequences. In particular sequences of P. neglectus and P. penetrans from

WA were most variable, with only 34-64% and 52-57% similarity with global populations of the same species, whereas ITS sequences of P. thornei had high correspondence (96-99%) with sequences identified overseas. These results might be interpreted as reflecting a more recent introduction of P. thornei to WA or that this species has a very low divergence.

Amplification and analysis of the D2-D3 amplicons from P. curvicauda allowed their comparison with those published by Hodda et al., (2014). The results obtained were very different from those published for P. quasitereoides, being up to 39% different when compared with the same parts of sequence provided by Hodda et al., (2014). Phylogenetic analysis of D2-D3 sequences clustered all the sequences of P. curvicauda studied in this study, separately from those of Hodda et al., (2014), which clustered together with sequences of P. teres. This is an important finding, in that now there are clearly two, possibly widely

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spread, endemic species in WA, in addition to the well-known P. neglectus, P. penetrans and

P. thornei.

The work undertaken in this study also confirmed that there could be mixed populations of RLNs in the same paddock, and emphasises that the application of successful management practices needs knowledge of which species are actually present.

7.3 Host preferences of P. curvicauda

With the previous conclusion in mind, the responses to infection of a range of plants to

P. curvicauda was assessed, using the reproduction factor, Rf (the ratio of final and initial nematode populations after ten weeks) as a measure of host response (at least in terms of susceptible or resistant). The Rf for this nematode species differed between host plants and also between cultivars of the same crop species. Cultivars of barley, sorghum, wheat and millet were good hosts with an Rf>1. The legumes (field pea, chickpea), brassicas (canola), pastures (biserrula, forage brassica, ryegrass) and the solanaceous plants, tomato and N. benthamiana, a native of Australia were intermediate hosts, and all supported some nematode replication, but without any increase in nematode population. These may be characterised as moderately resistant. Conversely, lupin cv. Jennabillup, triticale (AGG3959) and the genotypes tested of the pastures serradella, lucerne, persian clover and sub clover were all non-hosts of P. curvicauda. However, it is premature to characterise these species categorically as non-hosts, since these results were based on reproduction of nematodes under glasshouse conditions. Also, the experiments done did not extend to quantifying yield losses in the tested cultivars/germplasm with infection.

Crop plant genotypes normally exhibit four main types of response to pests and pathogens: resistance, susceptibility, tolerance and intolerance. Resistance and susceptibility relate to the ability of nematodes to reproduce in plants, while tolerance and intolerance relate

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to yield losses in the field. Cultivars identified as susceptible to P. curvicauda support reproduction of this species, and cultivars that are resistant can suppress or inhibit multiplication of nematodes in the field. For the same levels of nemtaode infestation, for tolerant genotypes yield losses in the field are comparatively lower than for plant genotypes decribed as intolerant for which yield losses are greater.

One comment about symptoms of P. curvicauda infestation of roots is that this species does not often cause any obvious brown lesions on host roots. If this is a general observation then that makes its presence more difficult to identify in the field. Another aspect which added to the difficulty of working with this nematode is that as yet it cannot be cultured in vitro, this is another difference from the other three RLN species under study, which could all be cultured and multiplied in vitro on carrot.

7.4 Biology of P. curvicauda

To determine the length of the life cycle of this particular species of Pratylenchus, J2 and

J3 stages were inoculated to susceptible wheat cv. Calingiri and studied until the same stages again were present in roots. This experiment showed that P. curvicauda took 35-42 d at 23°C in glasshouse conditions to complete one generation in wheat. This provides new information for P. curvicauda, although further studies along these lines targeting more host and different conditions would be useful.

Another feature of the biology of the interaction of P. curvicauda with host plants undertaken in this research was structural changes in host roots on nematode invasion. In general, at the level of study within the time available, histological changes observed in infested roots were similar irrespective of whether the host was of a resistant or susceptible genotype, at least for the genotypes of wheat and barley examined in this study. Root structures were examined by light and transmission electron microscopy 3 and 7 d after

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inoculation. There were clear changes at the cellular level, including cytoplasmic disturbance, disruption of affected cell walls and degraded cellular components, including mitochondria and nuclei, compared to images of control root cells which had not been inoculated with nematodes. One aspect that was new was that changes were evident in cells which were not next to the nematodes, including changes to the appearance of the nucleus.

Overall, the results obtained are very similar to those published (Vieira et al., 2017) and clearly show the massive damage caused to root cells and tissues in infested host roots. It is not surprising that, with this level of damage to cells of infested roots, there is a significant impact on crop yield from disturbing root functions and transport of nutrient and water through roots.

Studies were also undertaken to investigate attraction and penetration of P. curvicauda to host roots. Attraction to both resistant and susceptible genotypes was assessed. The results showed that nematodes migrated towards roots, penetrated and entered roots in the same numbers, irrespective of whether the host was classified as resistant or susceptible. Initially many nematodes were attracted to and entered test roots, but later penetration assay showed fewer nematodes after 14 d. It seems that nematode movement and penetration was not associated with crop genotype, that is, whether it was resistant or susceptible, unless the species was a non-host.

However, it was also evident that the pasture species assessed were less suitable host or non-hosts, and these could be good choice for growers to use in rotations to reduce P. curvicauda population levels in the field. In addition, there did not appear to be preferential penetration of the root tip zone. The finding that there were fewer nematodes in roots at 14 d compared to 7 d after inoculation could reflect the migratory nature of Pratylenchus, and the fact that as migratory endoparasites they are able to leave roots as well as enter them.

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In considering attraction of nematodes to roots, there are a number of underlying factors. Plant roots secrete many compounds into the rhizosphere. These include a range of metabolites and volatile compounds which can mobilise nutrients, attract beneficial microorganisms and modify the local environment. RLNs respond to these signals and migrate along gradients of metabolites, changes in pH and other signals to locate roots (Fosu-

Nyarko & Jones 2016). What signals RLNs respond to has not yet been studied, but it appears that these signals do not differentiate susceptible or resistant wheat genotypes, and that manifestation of resistance is at a different stage in the interaction. This interaction is different from a non-host response, in which attraction signals may be absent, or the nematode lacks effectors needed for invasion of that plant. This is an area of study that would benefit from biochemical or molecular studies to understand the molecular basis of interactions between RLNs and resistant or susceptible hosts.

Research on root-knot nematode suggested that PPNs release molecules in advance of root invasion to which epidermal cells of host may respond (Weerasinghe et al., 2005). At the site of root penetration further signalling occurs between the nematode and host cells, including a series of exchanges of nematode-associated molecules and host proteins (Fosu-Nyarko &

Jones 2016). The migration of nematodes in roots depends on gene products secreted from esophageal gland cells through the stylet hypodermis or amphids (Fig 7.1). Although the molecular basis of Pratylenchus-host interaction mechanisms is still at an early stage, there are more changes in gene expression in resistant cultivars than susceptible cultivars of the same crop (Backiyarani et al. 2014; Yu et al. 2015).

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Figure 7.1. Interaction of RLNs with host cells. Activities of effectors, proteolytic enzymes, and other proteins during migration and feeding by the nematode in host cells are indicated (Fosu-Nyarko & Jones 2016).

Invasion of host root tissues by Pratylenchus spp. triggers different signalling pathways during penetration and establishment in a host (Fosu-Nyarko & Jones 2016; Vieira et al.,

2017). The current state of knowledge of RLN-host cell interactions is shown in (Fig 7.1).

What this figure depicts is that RLNs secrete a range of cell-wall modifying proteins

(effectors), which presumably aid in penetration and migration of nematodes through root tissues, and a set of effectors which the nematodes use to avoid or evade host defence responses. However, they lack the effectors needed for feeding cell formation that are present in secretions of sedentary endoparasitic nematodes, and effectors which are common to all plant parasitic nematodes are likely to be involved in migration and suppression of host defences. These observations also help to confirm which effectors are needed for feeding site formation and which for migration and host defence suppression. Figure 7.1 also indicates that some effectors secreted by RLNs can influence neighbouring cells in advance of direct

115

interaction with the nematode, and the first evidence for this was found in this study in the ultrastructural studies, in which changes to nuclear cytoplasmic appearance were found.

Whether this represents transport of effectors from the nematode to neighbouring cells, or defence responses by the host cells, remains to be determined.

Looking to the future, study of the biology and host interactions of P. curvicauda has only just begun. Other RLNs are amenable to gene silencing using RNAi (Tan et al., 2013), and the same could be expected for P. curvicauda, but from a practical viewpoint, it is likely that classical screening methods will be used to screen new cereal germplasm for host responses for some time, combined with similar approaches for the other RLNs of economic importance.

7.5 Future recommendations

 Since P. curvicauda does not reproduce on carrot discs, it would be valuable to find a

suitable host to maintain and multiply P. curvicauda in vitro. This would enable large

numbers of monoxenic cultures to be generated for experimental work. However, the

crop cultivars found as suitable hosts for this nematode species could be used to

generate large numbers of nematodes for future experimental work.

 A more detailed survey is needed to map the presence and population density of P.

curvicauda in WA. It is also important to study mixed populations of P. curvicauda

and P. quasitereoides (if it can be found, or it can be established in pot cultures).

Moreover, it would be interesting to explore whether P. curvicauda has a competitive

advantage over P. quasitereoides (or other Pratylenchus spp. for that matter).

 This survey should include further analyses of the genetic variation that exists

between populations both at individual sites and at different locations, perhaps also

116

focusing on mitochondrial DNA sequences, and should also include more

morphological and morphometric data.

 The information on P. curvicauda should be included in the PredictaB service to

support crop producers and their advisors with the early managenemt practices against

this species.

 Next Generation Sequencing should be applied to study population variation in more

detail, and this could be extended to identifying effectors present and possible new

targets for chemical or genetic control.

 Although host responses and host preferences were studied in glasshouse

experiments, quantifying yield losses of the economic crops identified as hosts in the

field would be extremely valuable.

 More detailed ultrastructural studies may give a better understanding of host

responses of resistant and susceptible host genotypes.

Overall, the research undertaken in this thesis shows that there are still many challenges to working with root lesion nematodes and particularly with P. curvicauda, and that the results presented here provide new knowledge of this economically important crop pest.

117

Appendix

Table 1. ITS sequences of Pratylenchus spp. used for construction of phylogenetic tree available at NCBI

Name of Pratylenchus spp. Accession number Sequence Length Name of Pratylenchus spp. Accession number Sequence Length Pratylenchus agilis FJ712887.1 881 Pratylenchus crenatus LC030310.1 928 Pratylenchus agilis FJ712891.1 880 Pratylenchus crenatus LC030308.1 958 Pratylenchus agilis JQ039330.1 863 Pratylenchus fallax FJ712918.1 673 Pratylenchus agilis KC952982.1 882 Pratylenchus fallax FJ712919.1 673 Pratylenchus alleni JX081545.2 1080 Pratylenchus floridensis GQ988375.1 991 Pratylenchus araucensis FJ154951.1 1056 Pratylenchus floridensis GQ988376.1 991 Pratylenchus bhattii JN244271.1 788 Pratylenchus goodeyi FJ712922.1 784 Pratylenchus bhattii JN244272.1 788 Pratylenchus goodeyi KF840455.1 774 Pratylenchus bhattii JN244273.1 787 Pratylenchus goodeyi KF840456.1 784 Pratylenchus bolivianus FJ712892.1 952 Pratylenchus goodeyi KF856291.1 786 Pratylenchus bolivianus FR692329.1 920 Pratylenchus gutierrezi FJ712927.1 962 Pratylenchus bolivianus FR692328.1 914 Pratylenchus gutierrezi FJ712928.1 960 Pratylenchus bolivianus HM469447.1 949 Pratylenchus hippeastri FN554887.1 993 Pratylenchus brachyurus HQ662583.1 721 Pratylenchus hippeastri KJ001717.1 970 Pratylenchus brachyurus KF221216.1 919 Pratylenchus hippeastri FJ712932.1 964 Pratylenchus brachyurus KF534515.1 919 Pratylenchus jaehni FJ712937.1 997 Pratylenchus brachyurus KC538863.1 920 Pratylenchus jaehni FJ712940.1 997 Pratylenchus coffeae JN809832.1 953 Pratylenchus jaehni FJ712941.1 997 Pratylenchus coffeae JN809833.1 953 Pratylenchus japonicus KF452048.1 857 Pratylenchus coffeae JN809839.1 968 Pratylenchus japonicus KF452049.1 857 Pratylenchus coffeae KJ721977.1 1034 Pratylenchus kumamotoensis LC030314.1 899 Pratylenchus convallariae FJ712907.1 715 Pratylenchus kumamotoensis LC030312.1 892 Pratylenchus convallariae FJ712908.1 717 Pratylenchus kumamotoensis LC030317.1 899 Pratylenchus convallariae HM469448.1 722 Pratylenchus lentis AM933147.1 699 Pratylenchus crenatus LC030306.1 932 Pratylenchus lentis AM933148.1 698 118

Contd. Table ………

Name of Pratylenchus spp. Accession number Sequence Length Name of Pratylenchus spp. Accession number Sequence Length Pratylenchus lentis AM933154.1 703 Pratylenchus neglectus JX228136.1 750 Pratylenchus loosi FJ712942.1 997 Pratylenchus neglectus LC030323.1 871 Pratylenchus loosi JN091968.1 1030 Pratylenchus parafloridensis GQ988377.1 966 Pratylenchus loosi FJ712943.1 997 Pratylenchus parafloridensis GQ988378.1 961 Pratylenchus mediterraneus FJ712947.1 723 Pratylenchus penetrans FJ712958 717 Pratylenchus mediterraneus FR692306.1 946 Pratylenchus penetrans FJ712959 717 Pratylenchus mediterraneus FR692307.1 943 Pratylenchus penetrans FJ712962 712 Pratylenchus mediterraneus FR692310.1 755 Pratylenchus penetrans FJ712963 715 Pratylenchus neglectus FJ712953 680 Pratylenchus penetrans FJ712967 712 Pratylenchus neglectus FJ712954 681 Pratylenchus penetrans FJ712968 714 Pratylenchus neglectus FJ712955 683 Pratylenchus penetrans FJ712970 713 Pratylenchus neglectus FJ712956 683 Pratylenchus penetrans FJ712971 712 Pratylenchus neglectus FR692278 847 Pratylenchus penetrans FJ712982 720 Pratylenchus neglectus FR692279 849 Pratylenchus neglectus FR692282 849 Pratylenchus neglectus FR692280 845 Pratylenchus neglectus FR692283 853 Pratylenchus neglectus FR692281 845 Pratylenchus penetrans JX046943 782 Pratylenchus neglectus FR692285 853 Pratylenchus penetrans JX046944 779 Pratylenchus neglectus FR692288 852 Pratylenchus penetrans JX046945 776 Pratylenchus neglectus FR692289 850 Pratylenchus penetrans JX046946 777 Pratylenchus neglectus LC030327.1 833 Pratylenchus penetrans JX046947 779 Pratylenchus neglectus FJ712952 685 Pratylenchus penetrans JX046950 782 Pratylenchus neglectus FR692284 849 Pratylenchus penetrans JX046951 782 Pratylenchus neglectus FR692284.1 849 Pratylenchus penetrans JX046952 780 Pratylenchus neglectus FR692286 851 Pratylenchus penetrans JX046953 775 Pratylenchus neglectus FR692287 850 Pratylenchus penetrans FJ712957 711 Pratylenchus neglectus FR692287.1 850 Pratylenchus penetrans FJ712960 718

119

Contd. Table ……… Name of Pratylenchus spp. Accession number Sequence Length Name of Pratylenchus spp. Accession number Sequence Length Pratylenchus neglectus FR692290 854 Pratylenchus penetrans FJ712961 719 Pratylenchus penetrans FJ712964 713 Pratylenchus thornei FJ717821.1 672 Pratylenchus penetrans FJ712965 716 Pratylenchus thornei FR692299 828 Pratylenchus penetrans FJ712966 713 Pratylenchus thornei FR692299.1 828 Pratylenchus penetrans FJ712969 715 Pratylenchus thornei FR692300 826 Pratylenchus penetrans JX046948 783 Pratylenchus thornei FR692300.1 826 Pratylenchus penetrans JX046949 775 Pratylenchus thornei FR692301 830 Pratylenchus pseudocoffeae FR691856.1 942 Pratylenchus thornei FR692302 832 Pratylenchus pseudocoffeae LC030337.1 1090 Pratylenchus thornei FR692303 1001 Pratylenchus pseudocoffeae LC030338.1 1090 Pratylenchus thornei FR692304 974 Pratylenchus scribneri JX046932.1 889 Pratylenchus thornei FR692305 1001 Pratylenchus speijeri KF974726.1 1037 Pratylenchus thornei FR692305.1 1001 Pratylenchus speijeri KF974730.1 1026 Pratylenchus thorne FJ713006 800 Pratylenchus speijeri KF974737.1 1033 Pratylenchus thornei FJ713003 800 Pratylenchus speijeri KJ698683.1 1038 Pratylenchus thornei FJ713002 799 Pratylenchus speijeri KJ698684.1 1026 Pratylenchus thornei FJ713002.1 799 Pratylenchus penetrans FJ712984 720 Pratylenchus vulnus FR692320.1 915 Pratylenchus penetrans JX046942 780 Pratylenchus vulnus JQ003988.1 727 Pratylenchus scribneri JX046933.1 893 Pratylenchus vulnus JQ966892.1 915 Pratylenchus scribneri JX046934.1 957 Pratylenchus vulnus JX046959.1 765 Pratylenchus scribneri JX046935.1 949 Pratylenchus vulnus LC030340.1 860 Pratylenchus scribneri KM094195.1 883 Pratylenchus vulnus LC030341.1 863 Pratylenchus speijeri KF974719.1 1032 Pratylenchus zeae FJ643590.1 967 Pratylenchus speijeri KF974723.1 1028 Pratylenchus zeae JQ966893.1 904 Pratylenchus thornei FJ713005 799 Pratylenchus zeae KF765428.1 874 Pratylenchus thornei FJ713003.1 800 Pratylenchus zeae LC030346.1 847 Pratylenchus thornei FJ713004 799 Pratylenchus zeae LC030347.1 842 Meloidogyne hapla LC030360.1 713 Radopholus similis KF234227.1 854 120

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