Investigating Root-Knot and Cyst Parasitic Interactions through Transcriptomic Analyses of the Host and Parasite

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

By

Ellie Walsh

Graduate Program in Plant Pathology

The Ohio State University

2016

Dissertation Committee:

Professor Christopher G. Taylor, Advisor

Professor Thomas Mitchell

Professor Terry Niblack

Professor Margaret Redinbaugh

Copyrighted by

Ellie Kathleen Walsh

2016

Abstract

Plant-parasitic are a major threat to global agricultural production. Root-knot nematodes (RKN, Meloidogyne spp.) are arguably the biggest threat, capable of parasitizing virtually every crop. (SCN; glycines) has a narrow host range, but is the most destructive pathogen of a particularly important crop, soybean (Glycine max). RKN’s wide host range makes crop rotation often inadequate for management, and host resistance is unavailable in many crops. Effective resistance is available against SCN however populations have adapted to the most frequently used sources of resistance. RKN and SCN both induce elaborate feeding sites. In addition to being the sole source of nutrition, the feeding sites are the primary targets of nematode secretions to manipulate host cellular functions; consequently, they are very important interfaces of the interaction. The general aim of this research was to elucidate changes in the transcriptome underlying the successful interaction between these nematodes and their hosts. Although the use of RNA interference (RNAi) to knockdown nematode is actively being pursued as a new strategy for nematode control, little is known about the effects of general RNAi mechanisms during parasitism. As the suppression of RNAi has been characterized in other pathosystems, I hypothesized that parasitic nematodes may also be influencing these pathways. Tanscriptomic analysis of genes associated with RNAi machinery and target genes indicates that RNAi-regulated pathways are altered during the parasitic interaction. Using a silenced reporter , I found the disturbance to be specific to the nematode feeding site. Furthermore, disrupting these pathways with viral suppressors of RNAi renders the host more susceptible to nematode parasitism. Transcriptomic analysis indicates that this effect extends into later stages in parasitism, making the adult female stage of particular interest for further analyses. ii

I performed a transcriptomic analysis of adult female M. incognita to address the hypothesis that transcriptional patterns in this later stage of parasitism will reveal new candidate genes encoding that regulate the parasitic process, such as proteins that interact with RNAi among other plant pathways. Results from RNA-Seq analysis and reverse transcriptase PCR indicate that cell wall modifiers likely continue to play an important role in the parasitic interaction. Results from transcriptomic analysis including the putative secretome have highlighted new candidates for functional analysis to determine their role in the interaction. The later stage in parasitism is similarly of interest in SCN. SCN populations are adapting to the most commonly planted host resistance available in soybean, derived from Plant Introduction (PI) 88788 and “Peking.” The resistance response in PI 88788 appears to be longer-lasting than that in Peking, which impacts nematodes’ early development. Due to my focus on the adult female stage, I chose to investigate parasitism on PI 88788. I hypothesize that transcriptional differences between females from populations avirulent and virulent on PI 88788 may play a role in their adaptation to resistance. Results from this study indicate that the expression of an effector-like gene may have been lost in virulent populations, presumably allowing them to evade host detection and subsequent defense responses.

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Acknowledgements

The research detailed herein would not have been possible without a lot of guidance and support. I have been very fortunate to work in the Department of Plant

Pathology at The Ohio State University, which houses amazing faculty, fellow students, and staff. I would like to specifically acknowledge those individuals that have directly impacted this body of work. Therese Miller – the “mother of nematodes,” and Bob James

– the greenhouse keeper contributed significantly to the success of experiments. My

College of Wooster advisee, Allison Grenell, both executed experiments, and was instrumental to my own growth as a scientist and as a teacher; it is difficult to say who mentored whom. Numerous postdoctoral scientists have shared their time to teach me, namely Drs. Bryan Cassone, Gina Pengue, Phanikanth Turlapati, Yuhong Li, and Linjian

Jiang. I am also indebted for the tremendous efforts of Monica Lewandowski, Lynn

West, Ken Nanes, Niqui Beckrum, and Ramona Powell in handling graduate school logistics, likely more so than I will ever fully realize.

I would also like to thank all of my committee members for their support, unwavering confidence in me, and tremendous patience. Finally, I would like to express my gratitude and admiration for my major advisor, Dr. Christopher G. Taylor. His ability to convey his passion for research inspired me to pursue this degree and his guidance and relentless encouragement have allowed me to see it to completion.

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Vita

2008...... B.S. Plant Science, International Agriculture &

Rural Development, Cornell University

2010...... M.S.Ed. Science Education, CUNY Lehman

College

2008 – 2010...... Science Teacher, 7th grade, The Melrose School

2010 – 2012...... Graduate Fellow, Plant Molecular Biology &

Biotechnology, The Ohio State University

2012...... M.S. Plant Pathology, The Ohio State University

2013...... Graduate Teaching Assistant, Plant Disease

Diagnosis, The Ohio State University

2014...... Instructor, OSU International Diagnostics Short

Course Program, The Ohio State University

2015...... Instructor, Workshop entitled “Investigating Fungi

in the Environment,” University of Puerto

Rico, Mayaguez

2012 to present ...... Graduate Research Associate, Department of Plant

Pathology, The Ohio State University

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Publications

Testen AL, Walsh EK, Taylor CG, Miller SA, Lopez-Nicora H. (2014) First report of bloat nematode ( dipsaci) infecting garlic in Ohio. Plant Disease 98(6): 859- 859.

Veturi Y, Kump K, Walsh EK, Ott O, Poland J, Kolkman JM, Nelson RJ, Balint-Kurti P, Holland J, Wisser R. (2012) The longitudinal mixed linear model: an information- rich statistical technique for analyzing disease resistance data. Phytopathology 102(11):1016- 25.

Chung C, Poland JA, Kump K, Benson J, Longfellow JM, Walsh EK, Balint-Kurti PJ, Nelson RJ. (2011) Targeted discovery of quantitative trait loci for resistance to northern leaf blight and other diseases of maize. Theoretical and Applied Genetics 123(2):307- 326.

Chung C, Longfellow J, Walsh EK, Kerdieh Z, Esbroek GV, Balint-Kurti P, Nelson R. (2010) Resistance loci affecting distinct stages of fungal pathogenesis in maize: use of introgression lines for QTL mapping and characterization. BMC Plant Biology 10:103.

Field of Study

Major Field: Plant Pathology

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Table of Contents

Abstract ...... ii Acknowledgments ...... iv Vita ...... v Table of Contents ...... vii List of Tables ...... ix List of Figures ...... xii Chapter 1: Literature Review 1.1. Plant Parasitism in the Phylum Nematoda: Root-Knot and Soybean Cyst Nematode Pathogenesis ...... 1 1.2. Plant-Parasitic Nematodes: Effects on Agricultural Production ...... 6 1.3. Root-Knot Nematode Parasitism ...... 11 1.4. Host Resistance to Root-Knot Nematodes ...... 15 1.5. Soybean Cyst Nematode Parasitism ...... 16 1.6. Host Resistance to Soybean Cyst Nematodes ...... 18 1.7. Host Response to RKN & SCN Parasitism ...... 22 1.8. Role of RNA Silencing in Plant-Parasitic Nematode Research ...... 24 1.9. References ...... 28 Chapter 2: Root-Knot Nematode Parasitism Suppresses RNA Silencing 2.1. Abstract ...... 39 2.2. Introduction ...... 40 2.3. Materials & Methods ...... 42 2.4. Results ...... 45

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2.5. Discussion ...... 47 2.6. Acknowledgements ...... 50 2.7. References ...... 50 Chapter 3: The Mature Female Root-Knot Nematode Transcriptome 3.1. Abstract ...... 77 3.2. Introduction ...... 78 3.3 Materials & Methods...... 81 3.4. Results ...... 85 3.5. Discussion ...... 90 3.6. Acknowledgements ...... 94 3.7. References ...... 95 Chapter 4: The Mature Female Soybean Cyst Nematode Secretome & Adaptations to PI 88788 Resistance 4.1. Abstract ...... 122 4.2. Introduction ...... 123 4.3. Materials & Methods ...... 127 4.4. Results ...... 131 4.5. Discussion ...... 134 4.6. Acknowledgements ...... 139 4.6. References ...... 139 Chapter 5: Challenges, Implications, and Future Directions ...... 165 Bibliography ...... 172

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List of Tables

Table 1.1. Summary of method, target, and results of studies utilizing RNAi for plant- parasitic nematode control...... 37 Table 2.1. Disruption of RNAi-related gene expression observed during nematode

infection. Log2 fold changes in feeding site or infected treatment versus non-feeding site or uninfected control are presented with a false discovery rate of < 5% for giant cell (Morse et al., 2010) and syncytium studies (Szakasits et al., 2009) and P < 0.05 after Bonferroni correction for the gall study (Jammes et al., 2005). N/A are not differentially expressed at the significance cutoff, and black cells are not represented on the CATMA chip. Among significantly differentially expressed genes, the number of upregulated RNAi machinery genes deviate significantly from an expected ratio of 1:1 in laser- capture microdissected giant cells (Chi-square test, p < 0.0005), galls at both 14 and 21 days post infection in comparison to uninfected tissue (Chi-square test, p < 0.1), and in microaspirated syncytia (Chi-square test, p < 0.001). Among significantly differentially expressed miRNA-targeted genes in giant cells, the majority are upregulated (Chi-square test, p < 0.1). A disproportionate number of ta-siRNA-targeted genes are also upregulated in both giant cell and syncytium studies (Chi-square test, p < 0.05) ...... 66 Table 3.1. Primer sequences used for detection of Arabidopsis transcripts in M. incognita cDNA libraries ...... 101 Table 3.2. Primer sequences used for PCR validation of expression in M. incognita egg, J2, and adult female cDNA libraries ...... 102 Table 3.3. Top 15 most abundant transcripts expressed in adult M. incognita based on average coverage and paired-end reads mapped...... 107

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Table 3.4. Top 15 most abundant transcripts in adult M. incognita with putative secretion signal, based on average coverage and paired-end reads mapped...... 109 Table 3.5. RT-PCR validation of expression detected in RNA-Seq dataset from adult female M. incognita. RT-PCR relative expression results from egg and J2 cDNA libraries included for life stage comparisons...... 112 Table 3.6. Cell wall-degrading or modifying top BLASTx hits in female M. incognita transcriptome...... 115 Table 3.7. Expression (+) of 19 genes in adult females of 27 pioneer genes identified in a gland cell-specific M. incognita cDNA library (Huang et al., 2003). (-) indicates the absence of this accession in our dataset and therefore has no associated E-value (N/A).120 Table 3.8. Arabidopsis transcripts detected across four replicates of RNA-Seq analysis of M. incognita female cDNA libraries ...... 121 Table 4.1. Primer sequences used for detection of transcripts in female soybean cyst nematode cDNA libraries ...... 146 Table 4.2. Differentially expressed genes downregulated in populations virulent to PI 88788 relative to the avirulent soybean cyst nematode population, “Hancock,” grown on the susceptible cultivar, Lee 74...... 154 Table 4.3. Differentially expressed genes upregulated in soybean cyst nematode populations virulent to PI 88788 relative to the avirulent SCN population, “Hancock,” grown on the susceptible cultivar, Lee 74...... 156 Table 4.4. Genes uniquely downregulated in the virulent soybean cyst nematode populations (HG Type 2.5.7; Darke, Defiance, Erie, and Madison) relative to the avirulent population (HG Type 7; Hancock)...... 158 Table 4.5. Genes uniquely upregulated in the virulent soybean cyst nematode populations (HG Type 2.5.7; Darke, Defiance, Erie, and Madison) relative to the avirulent population (HG Type 7; Hancock)...... 159 Table 4.6. Genes uniquely downregulated in the virulent soybean cyst nematode populations HG Type 1.2.5.7 (Brown and Wood) and 1.2.3.5.7 (Knox and Pike)* relative to the avirulent population, HG Type 7 (Hancock)...... 160

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Table 4.7. Genes uniquely upregulated in virulent soybean cyst nematode populations (HG Type 1.2.5.7; Brown and Wood)* relative to the avirulent population (HG Type 7; Hancock) ...... 162 Table 4.8. Differentially expressed genes, upregulated in all populations of female soybean cyst nematodes grown on the resistant versus susceptible cultivar. Results do not include the HG Type 7 population (Hancock), as not enough females were produced for RNA isolation ...... 163

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List of Figures

Figure 2.1. expressing the TEV viral suppressor, HC-Pro, are more susceptible to M. incognita (RKN) infection. Data represent the mean ± s.e.m., n ≥ 19 plants; results were repeated in 2 additional replications (Wilcoxon, * p < 0.01) ...... 56 Figure 2.2. Relative absence of GUS expression in dsRNA:GUS two-week old seedlings (A) and restoration of GUS expression in progeny (B) of the dsRNA:GUS line crossed with P1/HC-Pro (line X-27-8, created by Mallory, 2001). The reciprocal cross also shows a restoration of GUS (not shown) ...... 57 Figure 2.3. Susceptibility of tobacco plants either homozygous or hemizygous for HC- Pro is enhanced. HC-Pro-expressing tobacco are significantly more susceptible to M. incognita than wild type in three independent experiments (data represent mean ± s.e.m., n ≥ 19, Wilcoxon * p < 0.01). Tobacco hemizygous for HC-Pro are also significantly more susceptible than the parent lacking HC-Pro (data represent mean ± s.e.m., n ≥ 15, Wilcoxon * p < 0.008) ...... 58 Figure 2.4. Two of three VSR-expressing Nicotiana benthamiana are more susceptible hosts to M. incognita. Data represent the mean ± s.e.m., n ≥ 24 plants; results were repeated in two additional replications with TCV CP and CRV P19 and one additional replication with PVA HC-Pro (Steel's Multiple Comparison Wilcoxon Tests, * p < 0.01) ...... 59 Figure 2.5. Silenced-reporter gene constructs are illustrated above. A) Expression cassette for dsRNA-silenced GUS. B) Individual expression cassettes for dsRNA- silenced GUS and SU-driven GUS. C) Individual expression cassettes for miRNA- silenced GUS and SU-driven GUS ...... 60

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Figure 2.6. Histochemical analysis of X-gluc stained dsRNA:GUS tobacco roots visualized with a dissecting microscope. Non-infected roots (A) and roots 14 dpi with J2 M. incognita wherein recovery of GUS expression is evident in knot tissue in close proximity to female M. incognita (B). Knot tissue was harvested for sectioning ...... 61 Figure 2.7. Restoration of dsRNA-silenced β-1, 4-endoglucanase (GUS) expression in nematode feeding sites. Sections taken at A) 4, B) 7, C) 14, and D) 21 days post inoculation with second-stage juvenile worms; GUS restoration evident in nematode (red*) giant cells (white*) (scale bar = 100 μm) ...... 62 Figure 2.8. Restoration of β-1, 4-endoglucanase (GUS) transcripts in nematode feeding sites, 7 dpi with juvenile M. incognita. RT-PCR products produced from uninfected and infected (knot) tissue; shown are three technical replications each representing three pooled plant samples. Note absence of GUS amplicon in uninfected tissue, and presence of GUS amplicon in knot tissue. Corresponding PP2A reference gene amplicon is presented along with no template control (NTC). Similar results were observed at 14 and 21dpi and repeated in three independent experiments ...... 63 Figure 2.9. RT-PCR products from non-inoculated tissue (lanes 1-3), noninfected tissue (lanes 4-6), knot tissue (lanes 7-9), and no template control (lanes 9). 1 kb β-1, 4- endoglucanase amplicon present in knot tissue at 7 (A), 14 (B), and 21dpi (C). Three technical replicates shown each from three pooled plant samples. PP2A 123bp amplicon from each corresponding sample in lower gel (100bp ladder, NEB) ...... 64 Figure 2.10. Restoration of silenced β-1, 4-endoglucanase (GUS) in nematode feeding sites. A) Early and late (inset) during M. incognita (*) infection in dsRNA-silenced SU- GUS whole roots. B) Cross-section of 21dpi M. incognita (*) feeding site in miRNA- silenced GUS roots ...... 65 Figure 3.1. Graphical depiction of the distribution of top tBLASTx hits of the transcriptome of the adult female on Arabidopsis. The majority of contigs returned no significant BLASTx hit (E value < 10-3), illustrated in the top pie graph. The lower pie graph illustrates the plant-parasitic nematode top BLASTx hits, representing 2% of the transcriptome assembly ...... 103

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Figure 3.2. Top twenty common terms (biological process) of putatively secreted proteins (blue) in adult female M. incognita transcriptome. Abundance of transcripts with these GO terms present in the whole transcriptome (red) ...... 104 Figure 3.3. Top twenty common gene ontology terms (cellular compartment) of putatively secreted proteins (blue) in female M. incognita transcriptome. Abundance of transcripts with these GO terms present in the whole transcriptome (red) ...... 105 Figure 3.4. Top twenty common gene ontology terms (molecular function) of putatively secreted proteins (blue) in adult female M. incognita transcriptome. Abundance of transcripts with these GO terms present in the whole transcriptome (red) ...... 106 Figure 3.5. Pairwise comparison results (E value < 10-10) of M. incognita 1,696 putatively secreted contig set with EST or cDNA databases from M. hapla, C. elegans, T. circumcincta, and D. viviparous...... 111 Figure 3.6. RT-PCR analysis of M. incognita cDNA template derived from eggs (1), J2 (2), and adult females (3). β-actin used for reference (A), pectate lyase (B), Cre-vit-6 (C), patatin (D), CM (E), VAP-1 (F), amphid-secreted (G), and eng-2 (H) (100bp ladder; NEB) ...... 113 Figure 3.7. RT-PCR analysis of M. incognita cDNA template derived from eggs (1), pre- parasitic J2 (2) and adult females (3). β-actin used for reference (A), chymotrypsin (B), 14-3-3b (C), VAP-2 (D), cathepsin-D (E) and expansin (F) (100bp ladder; NEB) ...... 114 Figure 3.8. RT-PCR analysis of M. incognita cDNA template derived from eggs (1), pre- parasitic J2 (2) and adult females (3).β-actin used for reference (A), transthyretin (B), gland 21 (C), calreticulin (D), and gland protein 23 (E) (100bp ladder; NEB)...118 Figure 3.9. RT-PCR results of select A. thaliana transcripts from female M. incognita cDNA libraries. Lanes 1-3: catalase (AT1G20620; 219bp, 476bp, and 872bp). Lane 4-5: TCTP (A T3G16640; 249bp and 640bp). Two independent replications in panels above, note negative result in second panel, lane 3. A transcript was considered “detected” when amplified in at least two independent cDNA libraries (100bp ladder; NEB) ...... 121 Figure 4.1. 32 soybean seedlings in individual cone-tainers were infested with infective juveniles of each soybean cyst nematode population in crocks containing either PI 88788

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or susceptible Lee 74 (A). Three days post infestation, acid fuchsin-stained juveniles can be observed (white arrow). The time point, 15 days post infestation (dpi), was chosen to optimize the number of females isolated prior to obvious melanization (18 and 24dpi) ...... 145 Figure 4.2. Soybean cyst nematode populations across Ohio were selected based on variation in geographic location, virulence, and HG Type. Populations include both HG Type 2 indicated in yellow, green, and blue, and non-HG Type 2 indicated in red and orange...... 147 Figure 4.3. Soybean cyst nematode populations vary in virulence to both the susceptible and resistant host; means from one HG Type assay are displayed. Color corresponds to population’s HG Type, red = HG Type 7, orange = 5.7, yellow = 2.5.7, green = 1.2.5.7, and blue = 1.2.3.5.7. Female Index (FI) thresholds are indicated with dotted lines (assays resulting in < 100 cysts on the susceptible checks are considered invalid)...... 148 Figure 4.4. Top twenty common gene ontology terms (cellular compartment) of putatively secreted proteins in female soybean cyst nematode transcriptome ...... 149 Figure 4.5. Top twenty common gene ontology terms (biological process) of putatively secreted proteins in female soybean cyst nematode transcriptome ...... 150 Figure 4.6. Top twenty common gene ontology terms (molecular function) of putatively secreted proteins in female soybean cyst nematode transcriptome ...... 151 Figure 4.7. Species distribution of top BLASTx hits of putatively secreted contigs in female soybean cyst nematode transcriptome ...... 152 Figure 4.8. Approximately ¼ of the female soybean cyst nematode transcriptome represents differentially expressed genes (DEG) between populations grown on a susceptible versus resistant line. Of the DEG, approximately 60% are unique to one population and the remaining DEGs are common to ≥2 populations. Sixty-seven DEGs are shared between all populations grown on the susceptible versus resistant line (green). Nine DEGs are common between the avirulent and virulent populations (orange)...... 153 Figure 4.9. A) Amplification of putative dorsal gland-secreted effector observed in lane 4, faint amplicon observed in lanes 3 and 5. B) Amplification of β-actin in corresponding

xv samples for reference. 1: Lucas (HG Type 7), 2: Brown (HG Type 1.2.5.7), 3: Defiance (HG Type 2.5.7), 4: Hancock (HG Type 7), and 5: Henry (HG Type 5.7) (100bp ladder, NEB; NTC = PCR no template control) ...... 155 Figure 4.10. Alignment of translated contig_4434 (subject) and the SNARE-like protein identified by Bekal et al. (2015), KM575849 (query). Figure adapted from blasp suite- 2sequences run on the NCBI webserver ...... 157

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CHAPTER 1: LITERATURE REVIEW

1.1. PLANT PARASITISM IN THE PHYLUM NEMATODA: ROOT-KNOT & SOYBEAN CYST NEMATODE PATHOGENESIS

Nematodes are the most abundant on the planet. As stated by the United States’ “father of nematology,” Nathan Cobb, “if all the matter in the universe except the nematodes were swept away, our world would still be dimly recognizable... we would find its mountains, hills, valleys, rivers, lakes and oceans represented by a film of nematodes.” Indeed, nematodes have been found to occupy extreme environments previously thought to be inhospitable to multicellular life. They successfully inhabit diverse settings, from the frigid Arctic (Holovachov, 2014) and Antarctic (Gradinger, 2001) to the high-pressure depths of the ocean floor, and even more than 3 kilometers down into the hot interior of the Earth’s crust (Borgonie et al., 2011). In soil and aquatic ecosystems, nematodes include plant feeders, bacterivores, fungivores, predators, and omnivores, and play an important role in nutrient cycling. Some nematodes have evolved as plant or parasites, and can cause significant damage to human health, as well as to livestock and agricultural production of food and fiber. The basic body plan of nematodes is well described by Decraemer and Hunt (2006) and more detailed morphological descriptions of root-knot nematodes (RKN) are provided in An Advanced Treatise on Meloidogyne (Eisenback, 1985). The outer layer of the nematode body consists of a cuticle (a body wall that is primarily comprised of collagen) that provides structure required for movement and protection from the external environment but is also semipermeable, allowing for the exchange of gases, water, and metabolites. Nematodes are included in the group Ecdysozoa due to their ability to periodically shed the cuticle through molting. Nematodes undergo four molts, 1 transitioning between four juvenile stages to an adult stage. Directly under the cuticle is the hypodermis, from which the cuticle is secreted, and somatic musculature running along dorsal and ventral sides, alternative contractions of which cause sinusoidal movement in the majority of nematodes. There are also dorsal and ventral nerve cords running the length of the body that converge at a nerve ring. Peripheral nerves connect the central nervous system to sensory protrusions and pores, allowing the nematode to sense and respond to its environment via chemical, mechanical, and thermal cues. Nematodes are pseudocoelomates, meaning they have a fluid-filled cavity between endoderm and mesodermal layers, which in lieu of a circulatory, respiratory, and skeletal system, is required for the diffusion of materials and provides turgor pressure necessary for body structure and movement. Interior to the psuedocoelom is the nematode’s digestive tract from its mouth (stoma and buccal cavity), esophagus (pharynx) and intestine, as well as a glandular or tubular excretory/secretory system and reproductive organs, leading to the rectum and anus and a cloaca or vulva respectively. The body plan of nematodes is relatively simple, yet abundant morphological, physiological, and ecological diversity exists within this group of animals, allowing them to occupy a wide array of ecological niches. The evolution of plant parasitism in the phylum Nematoda arose independently in at least three separate clades based on sequence analysis of small subunit ribosomal RNA (Blaxter, 2007; De Ley and Blaxter, 2002). A recent review by Quist et al. (2015) proposed four independent events: one clade, in the class , contains the most diverse group of plant-parasitic nematodes in the order , which encompasses both root-knot and soybean cyst nematode genera; a second clade also within Chromadorea contains the order Aphelenchida; and two additional clades in the class , orders and Dorylaimida. Each clade consists of free-living nematodes, particularly fungivores, as well as plant parasites. These independent events are evidence of convergent evolution of plant parasitism in nematodes. One feature common (but not exclusive) to all plant-parasitic nematodes is a hollow spear-like structure called a stylet. The stylet, used to puncture plant cells, ingest material, and

2 secrete small substances, may have originally evolved in fungivores, and then over independent events became useful for parasitizing plants (Quist et al. 2015). Another factor driving the evolutionary adaptation of plant parasitism in nematodes is horizontal gene transfer (HGT). The first strong indication of HGT events was the discovery of β-1,4-endoglucanases (enzymes that degrade cellulose and xyloglucan) produced in the esophageal glands of the cyst nematodes, and Heterodera glycines (Smant et al., 1998). Interestingly, cellulases are otherwise mostly absent in the animal kingdom. Cell wall modifying enzymes found by Smant et al. (1998) and by others in subsequent studies (Ledger et al., 2006; Rosso et al., 1999) share high sequence similarity with bacterial plant pathogens. Cell wall modifying enzymes are thought to aid in the nematode’s migration through host tissue. More comprehensive analyses indicate that over 3% of M. incognita protein-encoding genes originated from HGT events (Paganini et al., 2012). Other adaptations acquired via HGT will be discussed in detail in following sections as they pertain specifically to M. incognita and soybean cyst nematode (SCN) parasitism. Plant-parasitic nematodes are also often categorized based on their feeding strategy (Lambert and Bekal, 2002). Whereas most plant-parasitic nematodes are soil- borne root pathogens, some economically significant genera parasitize other plant tissues. Nematodes in the genus , for example, parasitize foliar tissue, spp. infect seeds, Ditylenchus spp. infect stem and bulb tissues, and Bursaphelenchus spp. infect the trunk (or stem) tissue of pine trees. Those nematodes that inhabit the soil ecosystem are further categorized according to their feeding strategy. Ectoparasitic nematodes remain on the outside of their host roots; they are motile and capable of feeding throughout their life cycle. This browsing-like feeding style has the benefit of allowing movement toward more suitable hosts but comes at the cost of vulnerability due to exposure to the soil environment. The genera Longidorus and Xiphinema in the order Dorylaimida exhibit this feeding behavior, and are able to serve as vectors of plant viruses as they move from one host to another. Migratory endoparasites also remain motile throughout their life cycle, but much of their time is spent migrating within host tissue, acquiring nutrition from and damaging plant cells along the way.

3 Sedentary nematodes lose their motility at some point during their life cycle; these (mostly female) nematodes exhibit either semi- or entirely endoparasitic feeding. Endoparasites have the ability to manipulate host cells into a permanent feeding site, which allows the females to focus much of their energy on reproduction. The semi- endoparasites, such as those in the genera Tylenchulus and , are attached to a permanent feeding site within a root, while the enlarged posterior portion of the body remains outside the root surrounded by soil. Sedentary endoparasites such as RKN and SCN are completely surrounded by host tissue when they initiate a feeding site, protected from the soil environment while they begin to feed. RKN and SCN both undergo their first molt within the egg, hatching as second- stage juveniles (J2). J2 are the infective stage, and migrate through soil toward host roots. The infective J2 predominantly enters roots near points of lateral root emergence and the actively dividing region behind root caps. They then migrate intercellularly through the cortex, circumvent the Casparian strip and situate their head into the root vasculature (Hussey, 1989). Although both RKN and SCN form permanent feeding sites, there are important differences between these nematodes’ life cycles. The RKN J2 punctures the cell walls of procambial cells near xylem sieve elements (Hoth et al., 2008) leaving the cells’ plasma membrane intact. Through the secretion of a variety of peptides, the nematode elicits mitosis followed by aborted cytokinesis such that the cells do not divide (Caillaud et al., 2008; Jones and Payne, 1978) and become “giant cells.” The cell walls of giant cells thicken and become highly invaginated, and their cytoplasm becomes dense with an abundance of small vacuoles, Golgi bodies, mitochondria, and endoplasmic reticulum. A unique organelle also forms within the cells in close association with the nematode’s stylet; this structure is referred to as a feeding tube, presumably to aid in the ingestion of cellular contents (Hussey and Mims, 1991; Rumpenhorst, 1984). When mature, giant cells are approximately 400 times the size of other root vascular cells and contain more than 100 polyploid nuclei (likely a result of endoreduplication) (Caillaud et al., 2008). A successful feeding site is composed of approximately 5 to 7 giant cells. The resulting giant cells act as a nutrient sink in the host, diverting nutrients to the feeding nematode from the rest of the plant

4 (Hofmann and Grundler, 2007). RKN unsuccessful at forming a feeding site due to unfavorable conditions (e.g. an abundance of competition or pruning stress) mature into adult males (Davide and Triantaphyllou, 1967; Snyder et al., 2006) and exit the root. M. incognita males have no role in reproduction as this species reproduces via mitotic parthenogenesis. Roughly thirty days after invading the root, RKN females have laid 150 to 300 eggs into a protective gelatinous matrix produced from rectal gland secretions (Bird, 1972). The eggshells themselves are also quite robust, composed of roughly 50% protein, 30% chitin, and lipids (Bird and McClure, 1976). RKN J2 develop and hatch at random under conducive environmental conditions, without the requirement of any specific external cue (Perry, 1997). Rather than forming a feeding site comprised of giant cells, SCN juveniles initiate the formation of a syncytium while developing into mature females. Males also begin to feed but, in contrast with RKN, evidence suggests that their sex determination has a significant genetic component, such that approximately 50% of developing SCN will become males, although the observed surviving proportions may differ (Colgrove and Niblack, 2005). Males regain their somatic musculature and exit the root to fertilize the exposed mature female (Sobczak et al., 1997). Similar to RKN-induced giant cells, the syncytium is induced by secretions of a variety of nematode peptides; however, SCN initiates the feeding site with a single root cell, which within 24 hours begins to incorporate surrounding cells along the vasculature via the dissolution of cell walls and protoplast fusion (Hussey and Grundler, 1998). The resulting syncytium has a dense cytoplasm, an abundance of small vacuoles, ribosomes, mitochondria, and feeding tube structures, and like giant cells becomes a nutrient sink. Within 21 to 30 days post- infection, the majority of the posterior female body has ruptured through to the exterior of the root, exposed for fertilization. Depending on conditions, the female’s body can fill with an average of 200 and upwards of 600 eggs (Sipes et al., 1992), 1/3 of which are deposited out of the female’s vulva into a gelatinous matrix (Ishibashi et al., 1973). As the female SCN dies, its cuticle melanizes and body becomes a cyst protecting the eggs within. Only a portion of eggs hatch at random without any external cue (Yen et al., 1995). 5

1.2. PLANT-PARASITIC NEMATODES: EFFECTS ON AGRICULTURAL PRODUCTION

Damage caused by plant-parasitic nematodes to agricultural production worldwide has been estimated at $80 to $125 billion every year (Nicol et al., 2011). Estimates of economic loss vary widely at this scale because nematode damage often causes non-specific symptoms and therefore is undetected and under-reported (Wang et al., 2003). Characteristic symptoms on RKN-infected root systems may be evident as root knots (galls) and cysts may be visualized on soybean roots infected with SCN. Above-ground symptoms however, often resemble those of a plant experiencing poor nutrient and water uptake such as chlorosis and stunting, symptoms which can be the result of several other factors. Depending on the conditions, even these nondescript symptoms may be virtually absent, although significant decreases in yield are still observed (Niblack, 2004) Both RKN and SCN damage are often also accompanied by secondary infections. The penetration and migration through root tissue provides other soil microbes additional access to root tissues, but secondary infections have mostly been found to benefit from nematode-directed physiological changes in the host (Karssen et al., 2013) and are dependent on a variety of other factors including host cultivar, isolate of the organism present, and environmental conditions (Webster, 1985). RKN is the most economically damaging plant-parasitic nematode on the planet (Trudgill and Blok, 2001). This is likely due to their worldwide distribution and ability to parasitize more than 2,000 plant species (Sasser 1980). More than 90 different species of RKN (Meloidogyne spp.) have been described to date; however, only four of the species cause the majority of the agricultural damage: M. arenaria, M. incognita, and M. javanica are present in areas where soils do not freeze, and M. hapla is present in temperate regions around the globe (Moens et al., 2009). M. incognita is found in every country around the world, making it the most widely distributed, and with its extensive

6 host range, arguably the most destructive pathogen in agricultural production (Trudgill and Block, 2001). In the United States, SCN is responsible for the greatest losses of soybean yield due to disease (Wrather et al., 2010), and economic losses have been estimated at $1.1 billion annually (Schmitt, 2004). Unlike RKN, SCN (Heterodera glycines Ichinohe) is a relatively host-specific parasite of soybean (Glycine max (L.) Merr.), although SCN is able to infect a broad range of other members of the family and more than 66 different weed species (Riggs, 1992), a matter that is relevant to management decisions. SCN is present in almost all of the soybean-production regions around the world. They were first detected in Ohio in 1987 (Riedel et al. 1988), and are now known to inhabit at least 68 of Ohio’s 88 counties (Dorrance et al. 2012). In addition to the damage incurred by SCN alone, other pathogen pressures have been found to compound resulting yield losses. Sudden death syndrome (SDS) caused by the fungal pathogen Fusarium virguliforme (Aoki, O’Donell, Homma, and Lattanzi) for instance, can be mildly aggressive, but in the presence of SCN, causes severe SDS symptoms (Westphal et al., 2014). Although synergism is not always observed and likely dependent on several factors including environmental conditions, soybean cultivar, level of inoculum, and population or isolate (Gao et al., 2006). Another potential disease complex has been found with charcoal rot caused by the fungal pathogen, Macrophomina phaseolina (Tassi) Goidanich (Mihail, 1992). M. phaseolina, like F. virguliforme, has been found to co-occur with SCN. Both SDS and charcoal rot individually have been estimated to be in the top five diseases causing soybean yield loss (Wrather et al., 2010). A potential association with the above-ground pest, the soybean aphid Aphis glycines Matsumura, was implicated (McCarville et al., 2014), but others found their effects to be independent of SCN (Hong et al., 2011). These findings illustrate that disease interactions in the field are difficult to determine, as a number of factors affect both agents and the complex. Potential interactions with other soybean pathogens, in addition to compounding yield loss, can also complicate the implementation of effective control strategies.

7 Multiple control strategies are integrated by farmers in order to maximize yield and profit, with a focus on whichever pests or pathogens are most damaging in a given area on a particular crop. RKN is a focus pest in many cropping situations (Karssen et al. 2013). Host resistance is the most economical strategy employed against RKN (reviewed in section 1.5.); however, effective resistant host options are not available for all crops. Many high-value crops (e.g., strawberry, citrus, and banana) warrant the use of chemical applications, including the use of fumigant and non-fumigant nematicides, but their adverse effects on the environment and health hazards have caused their use to be restricted and many are no longer produced in the United States (Haydock et al., 2013). Nonetheless, nematicides continue to be an important component of nematode management; of the estimated $1 billion worth of nematicides produced every year, approximately half are used for control of RKN mostly in vegetable and potato crops (Haydock et al. 2013). A variety of cultural practices are also employed to reduce nematode damage. For example, farmers in areas with sufficient sunlight employ soil solarization, a labor-intensive practice that is only reasonable to reduce nematode populations for high-value crops. The use of biological control agents and biopesticides (organisms and products derived from such organisms that are used to reduce pathogen populations and/or damage) has been increasingly investigated for the control of plant- parasitic nematodes, particularly as the use of nematicides has become more restricted. Significant advances have been made in the development of this strategy, yet more research is needed to tackle its variable efficacy, particularly under field conditions (Hallmann et al., 2009). Crop rotation is another management strategy, and while ineffective for many RKN species that have a wide host range that includes many weed species, crop rotations with non-host graminaceous crops can sometimes be effectively employed (Karssen et al., 2013). Unlike RKN, for which control strategies vary as widely as their host range, management of SCN revolves primarily around the use of crop rotation and host resistance. Crop rotation is cited as the most effective control strategy currently available for managing SCN, as it is the only strategy that reliably reduces associated yield losses (Niblack, 2005). Suggested rotation sequences include non-host crops (e.g., corn, barley,

8 canola) and resistant cultivars, rotating between different sources of resistance, and susceptible cultivars. More specific rotation recommendations have also been made based on the level and virulence phenotype of SCN populations detected in particular fields (Niblack and Chen, 2004). Yield loss estimates based on initial population levels are often confounded by other factors encountered in the field (Donald et al., 2006), making management decisions based solely on potential yield loss less reliable than decisions based on managing the SCN populations themselves. While decreasing soybean production with too many non-soybean plantings (commonly corn in the Midwest) is not always palatable to farmers, rotation between soybean cultivars with different sources of resistance can still be useful in managing SCN (described in section 1.6). Leaving fields fallow is another management strategy for plant-parasitic nematodes; however, it is rarely economically viable, and may be ineffective for nematode species able to reproduce on weeds. Complicating the effectiveness of this practice further for SCN is the longevity of their encysted eggs, which were found to produce infective J2 after seven to nine years without a host (Inagaki and Tsutsumi, 1971). The use of nematicides is generally successful at reducing nematode populations but rarely cost-effective in commodity crops such as soybean. The development of biological controls for SCN management is also being investigated, with challenges similar to their use in managing RKN that will need to be overcome before their use becomes prevalent. Advances for the management of RKN and SCN by means of genetic engineering have also been made. Transgenic approaches have been designed to interfere with nematode parasitism both directly and indirectly (these approaches are reviewed by McCarter, 2009). An example of direct interference is the expression of proteins that are toxic to the nematode. For example, cystatins (cysteine protease inhibitors) are thought to confer their activity by interfering with enzymes in the nematode’s gut that are essential to digestion. Cystatins have been cloned into the model plant Arabidopsis, resulting in a reduction of female size and fecundity in RKN and beet cyst nematode () females (Urwin et al., 1997) and have also proven effective in potato against the () (Urwin et al., 2001). The use of Crystal (Cry) proteins (δ-endotoxins produced by the soil bacterium Bacillus thuringiensis) are

9 also being explored, as these Cry proteins become activated in, bind to, and damage the midgut of a wide range of insects (reviewed in Palma et al., 2014). Some of these Cry proteins are toxic to a diverse set of species in the phylum Nematoda (Wei et al., 2003) including RKN (Li et al., 2007). Another direct approach being investigated to hinder nematode parasitism is the use of small RNA molecules targeting nematode transcripts, a method referred to as RNA interference (RNAi; reviewed in section 1.8). Rather than targeting the nematode itself, indirect approaches include those methods directed at altering or impeding the development of nematode feeding sites. While expressing a nematode-specific toxin has potential and has proven successful in its wide-scale use for insect control (e.g., Bt toxin-expressing crops), it has also been noted that this method can exert a strong selective pressure for resistance to the toxin in the pest being managed (Tabashnik et al., 2008). This and other pitfalls, such as downregulation of the constitutive promoters used for the expression of these toxins during nematode infection (Goddijn et al., 1993), prompted some research groups to examine the use of nematode-induced regulatory elements and targeting effects to the feeding site rather than the nematode directly. Work done with the root-specific TobRB7 promoter provides such an example, wherein expression was found to be upregulated specifically within giant cells and therefore provides an attractive target and tool to disrupt infection (Opperman et al., 1994). These types of promoter analyses have broadened transgenic crops’ potential in expressing desired traits, including combating various pathogens (reviewed in Potenza et al., 2004). Developments in our understanding of the molecular aspects of nematode feeding sites has expanded the possibilities that genetic engineering approaches have to offer; our current understanding of RKN and SCN feeding sites is discussed in Sections 1.3 and 1.4, respectively. It is evident that RKN and SCN represent a formidable hindrance to agricultural production around the world. No single management strategy has proven successful at eliminating these parasites, nor should a single effective measure be sought. Certainly a combination of management practices will continue to be the best way to reduce damage caused by these plant-parasitic nematodes and optimize yields and profits. While the use of chemical nematicides becomes more restricted, more sustainable methods such as

10 biological controls and diverse sources of host resistance require more research to increase their efficacies and broaden their application. In order to improve our options for nematode control and keep pace with adapting populations, there is still much to be gleaned from these parasites’ basic biology. The general aim of this thesis work was to elucidate mechanisms underlying a successful parasitic interaction between these nematodes and their hosts.

1.3. ROOT-KNOT NEMATODE PARASITISM

Successful RKN parasitism relies upon the ability to locate a suitable host root, migrate between root cells, and “reprogram” host cells into a feeding site. The reprogrammed cells must then be capable of providing all of the female’s nutritional requirements through reproduction. These major aspects of successful parasitism beg the question: how do RKN so drastically modify root cell morphology and physiology to their benefit while simultaneously evading host defenses? One approach to this question is to examine what compounds (or more specifically, effectors) are produced and secreted by the nematode throughout the course of infection. The functions of RKN effectors characterized to date along with some of their host targets were recently reviewed (Davis et al., 2008; Rosso et al., 2012 Truong et al., 2015). Prior to root invasion, RKN J2 are known to utilize chemotaxis to locate host roots in the soil environment, following gradients of carbon dioxide, for example (Pline and Dusenbery, 1987). In addition to perceiving signals from their environment, the nematodes’ anterior chemosensory receptors, amphids, are also actively secreting substances during migration through the soil and after entry into the host tissue (reviewed in Perry, 1996; Stewart et al., 1993). Some of these amphidial secretions may also play a role in chemoreception (Perry, 1996). Upon invasion of the host root, RKN rely upon an assortment of plant cell wall-modifying enzymes secreted from the nematode’s subventral esophageal glands (Ledger, 2006; Mitreva-Dautova 2006; reviewed in Vanholme, 2004). These enzymes, including cellulases and pectinases, are likely required

11 for the J2 to penetrate the host root and migrate intercellularly before initiating the formation of a feeding site. As discussed in Section 1.1, there are multiple lines of evidence that many of these enzymes were acquired through HGT (Ledger, 2006; Scholl and Bird, 2005). The plant’s endogenous cell wall modifying enzymes are also affected during the interaction. Through promoter analysis, Mitchum et al. (2004) found the expression of Arabidopsis gene Cel1 (a β-1,4-endoglucanase) to be induced during RKN infection, presumably due to its predicted role in the cell wall remodeling that occurs during giant cell formation. The authors did not observe similar induction during cyst nematode parasitism (beet cyst, Heterodera schachtii Schmidt and tobacco cyst, Globodera tabacum Lownsbery and Lownsbery), an example of another difference between the formation of giant cells and syncytia. After successfully breaching the physical barriers imposed by plant cell walls, RKN must also contend with other host defenses during invasion of the root. Throughout the early stages of parasitism, nematodes encounter the plant’s general defense response, which includes reactive oxygen species (ROS) like superoxide and hydrogen peroxide (Baker and Orlandi, 1995). Scavengers of ROS such as superoxide dismutase and ascorbate peroxidase are secreted by RKN, protecting the J2 as it migrates between cells (Molinari and Miacola, 1997). Because the nematode is reliant upon a functional feeding site through to reproduction, the evasion of host defenses remains important during this final stage of parasitism. The adult female has been found to produce catalase, another ROS scavenger, which perhaps plays a role against defense responses and/or detoxifying the metabolically active giant cells (Molinari and Miacola, 1997). Adult females were also shown to express a glutathione S-transferase, as determined by subtractive hybridization with J2, which may similarly confer protection from oxidative stress to the feeding nematode (Dubreuil et al., 2007). Calreticulin is another expressed protein, which is secreted and has been found to accumulate in cell walls (Jaubert, 2005). Calreticulin is known to dampen defenses in other animal eukaryotic parasites (Kasper, 2001; Suchitra and Joshi, 2005). The role of secreted calreticulin may have multiple functions as studies

12 have also indicated its potential influence on calcium signaling and cell cycle regulation (Borisjuk, 1998; Ghiran, 2003). Because the majority of virulence factors are produced and secreted from the esophageal glands of both RKN and SCN, research has focused on the isolation of these cells. Some investigators have induced secretions by stimulating the nematode’s metacorpus with the chemical resorcinol, and analyzed the secreted contents with mass spectrometry (Bellafiore et al., 2008; Jaubert et al., 2002). Another approach is microaspiration to harvest RNA specific to the esophageal gland cells, and analysis of the transcripts for candidate effectors (Davis et al., 2004; Davis et al., 2008). A technique has also been devised for the isolation of whole esophageal gland cells that may accelerate similar analyses in the future (Maier et al., 2013). Progress has been made in the identification and characterization of RKN and SCN effectors, as around 100 RKN effectors have been cloned (Truong et al., 2015). Only a few effectors appear to be common between RKN and SCN; chorismate mutase is one such example. Chorismate mutase (CM) is an enzyme in plants’ (and microbes’) shikimate pathway and has not been found in other members of the animal kingdom outside of plant-parasitic nematodes. Huang et al. (2005) characterized two CM effectors primarily expressed during early stages of parasitism, MI-CM-1 and MI-CM-2, and confirmed their CM activity via complementation experiments in Escherichia coli (Migula) Castellani & Chalmers. These CMs provide another potential example of a trait acquired via HGT. Chorismate-derived compounds have diverse roles in the cell; thus, whether the secretion of these proteins aids in evading defense versus altering the physiology of these cells will require further investigation. Some small secreted peptides have been found that mimic plant proteins. Expression analysis was used to identify 16D10, which encodes a small peptide with some homology to CLAVATA3 (CLV3) of Arabidopsis thaliana (Linnaeus) Heynhold (involved in the differentiation of meristematic cells). Expression of 16D10 in planta stimulated root growth and yeast two-hybrid tests showed a direct interaction with two of the host’s SCARECROW-like (SCL) transcription factors (Huang et al., 2006). When expressed in planta, 16D10 did not restore the phenotype in the clv3-1 mutant line,

13 suggesting that this homology does not indicate identical functions. Results indicate that 16D10 imparts its virulence function by eliciting SCL-mediated expression responses in the host. The development of resistance-breaking strains of nematodes (host resistance discussed in Section 1.5) has propelled studies into nematode avirulence factors as well. Near-isogenic lines of RKN were created, differing in their virulence to the resistance gene, Mi-1 (Jarquin-Barberena et al., 1991). One avirulence factor identified from this resource was MAP-1, an amphid-secreted protein, which is presumably secreted into the intercellular space and thus detected by the host early during the infection (Semblat et al., 2001). Avirulence genes expressed in RKN amphids and esophageal glands have been found, but aside from their elicitation of plant defense, many of their functions are still not entirely known (Neveu et al., 2003; Semblat et al., 2001). A cathepsin L protease, for example, found to be an avirulence factor in the presence of the Mi resistance gene, is expressed in the nematode intestine (Neveu et al., 2003). These examples expose an interesting facet that not all parasitism factors are secreted from the esophageal glands, but the amphids, digestive tract, and cuticle as well. Further work in this area will be necessary to better understand host resistance and the mechanisms underlying compatible versus incompatible reactions. The classical tradeoff hypothesis in which virulence comes at the cost of other life history traits may not adequately describe (or perhaps more likely is undetectable), as a study comparing avirulent and virulent near isogenic lines of RKN, found no associated tradeoffs (Castagnone-Sereno et al., 2015). While significant advances have been made in the identification and characterization of RKN effectors and other genes involved in parasitism, the identity and function of the majority of such proteins remains elusive. These proteins are currently characterized as “pioneers” because they have no known homologs, and thus have no revealing hints to their potential function(s). The ability to perform functional analyses for these proteins is improving with the use of RNAi (discussed in section 1.8). The objective of this thesis work was to determine the transcriptional profile of adult female RKN (specifically M. incognita) in order to examine the secretome during the latest stage of infection.

14

1.4. SOYBEAN CYST NEMATODE PARASITISM

SCN induces the formation of a single syncytium from which to feed. This feeding site has many properties and functions similar to those of giant cells, yet as would be expected for distantly related nematodes, there are also several differences. Research on SCN parasitism has focused on the host response to infection, identifying genes involved in parasitism in the nematode, and deciphering mechanisms of resistance in soybean (discussed in section 1.6). As more genes involved in parasitism have been identified, the majority of putative effectors appear to be dissimilar between RKN and SCN. One such novel effector identified in SCN, Hg19C07, was discovered in the cyst nematode’s dorsal esophageal gland. To better determine its interaction in Arabidopsis, researchers worked with its ortholog in beet cyst nematode, Hs19C07. Work by Lee et al. (2010) showed that Hs19C07 interacts with the auxin influx transporter LAX3 and that both LAX3 and a polygalactoronase (a LAX3-induced cell wall-modifier) are both upregulated in and around the developing syncytium. Additionally, lax3 mutants (in combination with other auxin influx mutants such as aux1) are more resistant to nematode infection (Lee et al., 2011). Although auxin was previously found to be important for syncytium development (Goverse et al., 2000; Grunewald et al., 2009), the involvement of this nematode effector was an important discovery in how SCN manipulate the host’s phytohormone balance. Plant signal peptide mimics have been discovered amongst SCNs’ effector repertoire. HgCLE1 (Hg-SYV46) and HgCLE2, secretory proteins with homology to CLV3 of Arabidopsis (similar to RKN’s 16D10) were characterized from SCN (Wang et al. 2005; Wang et al., 2010). Unlike 16D10, when cloned, HgCLE1 and HgCLE2 were able to complement the phenotype of the clv3-1 mutant. Thus, the specific actions of these effectors and 16D10 are likely different, wherein HgCLE1 and HgCLE2 are functional ligand mimics important to the developmental reprogramming and maintenance of syncytia.

15 As mentioned, chorismate mutase is an effector secreted by both RKN and SCN. The effector, Hg-CM-1, was confirmed to have chorismate mutase activity in planta and is predicted to suppress shikimate chorismate-derived compounds involved in defense (Bekal et al., 2003). Two alleles were identified, wherein Hg-cm-1A (or a closely linked gene) is selected for on PI 88788, but not on “Hartwig” (which contains resistance derived from PI 437654 and Peking (Anand, 1992)), PI 90763, or the susceptible “Lee 74” (Lambert et al., 2005). Selection of alleles may indicate this effector’s relevance to the adaptation to PI 88788 resistance. Much like RKN effectors, many SCN effectors lack homology to known proteins and so their potential function remains unknown. The objective of this thesis work was to determine the transcriptional profile of adult female SCN in order to examine the secretome during the last stage of infection.

1.5. HOST RESISTANCE TO ROOT-KNOT NEMATODES

Host resistance has played an important role in the management of plant-parasitic nematodes, and has been pursued as an attractive alternative to the use of pesticides (Roberts et al., 1986). Davies and Elling (2015) recently reviewed the current status and future outlook of host resistance to plant-parasitic nematodes. As discussed in Section 1.2, the withdrawal of nematicides from the market has highlighted the increasing importance of utilizing host resistance for nematode control. Several single resistance genes (R genes) as well as quantitative trait loci (QTL – multiple genes underlying the trait) have been identified and characterized to date. The first major R gene conferring resistance to RKN was introgressed from a wild species of , Lycopersicum peruvianum Mill, into the domesticated species L. esculentum (now Solanum lycopersicum L.) in the 1940’s (Smith, 1944). This R gene (Mi-1) confers resistance to three of the top four economically devastating root-knot nematodes, M. incognita, M. arenaria, and M. javanica, and is now extensively used around the world in several cultivars as well as a source of resistance in rootstocks. Mi-1

16 (specifically Mi-1.2) contains a coil-coil, nucleotide-binding, and leucine-rich repeat domain, characteristic of a class of R genes (CC-NBS-LRR), and leads to a hypersensitive response (HR), leading to localized cell death (Milligan et al., 1998). Melillo et al. (2006) further characterized this response, noting the burst of reactive oxygen species (ROS) indicative of HR and cell death within 24 hours post infection. Without a live feeding site, RKN mature into adult males. Interestingly, the Mi-1 gene was found to confer resistance to pests belonging to a different phylum, the potato aphid, Macrosiphum euphorbiae (Thomas) (Rossi et al., 1998; Vos et al., 1998), and white fly, Bemisia tabaci (Gennadius) (Nombela et al., 2003). Likely due to its widespread use, Mi- 1 resistance-breaking RKN have developed, leading to identification of additional R genes from L. peruvianum, Mi-3 and Mi-9, which are effective against some of these newly virulent populations (Yaghoobi et al., 2005). The most recently cloned and characterized R gene, Ma, was isolated from plum (Prunus cerasifera Ehrhart) (Claverie et al., 2004). Ma is in the toll/interleukin1 receptor nucleotide-binding site leucine-rich repeat (TNL) class of R genes, with an unusually large and highly polymorphic post-LRR domain (1,088 amino acids). Through complementation testing with A. rhizogenes-transformed roots, Ma conferred broad- spectrum resistance, with efficacy against a species unaffected by Mi-1 (M. enterolobii) (Claverie et al., 2011). Also, unlike Mi-1 (Philis and Vakis, 1977; Tzortzakakis and Gowen, 1996), Ma remains effective at higher temperatures and under high inoculum pressure (Claverie et al., 2001). These characteristics make Ma an attractive source of resistance. The majority of host resistance responses to RKN have been found to involve HR (reviewed in Davies and Elling, 2015), with the one exception of the Rk gene in cowpea, which allows M. incognita to invade and initiate giant cells for up to two weeks before the feeding site degenerates (Das et al., 2008). Whereas most RKN resistance responses are HR-dependent, the SCN resistance response conferred by PI 88788 tends to be delayed, allowing the nematode to establish a feeding site before female development and reproduction is hindered (reviewed in Davies and Elling, 2015).

17

1.6. HOST RESISTANCE TO SOYBEAN CYST NEMATODES

Resistance to SCN was sought immediately after their initial discovery in the United States in the 1950’s (Ross and Brim, 1957; Winstead et al., 1955). Incorporating resistance into soybean cultivars imparted an enormous benefit to soybean production. Many of the first cultivars developed did not compete well with traditionally grown varieties in non-infested fields, but the development of resistant cultivars with better yield potentials ensued. The introduction of the cultivar Forrest for example, prevented an estimated $400 million in yield lost to United States soybean farmers (Bradley and Duffy, 1982 cited in Miller, 1986). Several genes are indicated in host resistance to SCN. Inheritance studies have uncovered the existence of multiple loci, two independently inherited recessive loci, rhg2 and rhg3 (Caldwell et al., 1960), and dominant loci, Rhg1, Rhg4 (Matson and Williams, 1965; Meksem et al., 2001), and Rhg5 (Rao-Arelli et al., 1992). Upon the examination of QTL for resistance to SCN, variability exists in the level of resistance conferred (reviewed in Concibido et al., 2004). A report initially investigated a canonical R gene in the region of the Rhg1 QTL (Srour et al., 2012), however studies examining the LRR-kinase in this region found it had no detectable effect on SCN resistance and was eventually confirmed to not be the factor underling Rhg1 (Cook et al., 2012; Melito et al., 2010). Rhg1 resistance was later found to be conferred by 10 tandem copies of a 31kb region (versus one copy in susceptible cultivars such as Williams 82; and also has corresponding higher transcript levels) containing an amino acid transporter (Glyma18g02580), an α-soluble N- ethylmaleimide-sensitive factor attachment protein (α-SNAP), and an unknown protein containing a wound-inducible domain (WI12). These three dissimilar genes only confer resistance when overexpressed as a set (not when overexpressed individually) while their individual silencing decreases resistance (Cook et al., 2012). A look at the epigenetic markings also revealed a correlation of hypermethylation of Rhg1 genes in high copy,

18 resistant lines (Cook et al., 2014). Action of Rhg1 is thought to disrupt the maintenance in addition to the initiation of the syncytium. Resistance conferred by Rhg4 was also initially thought to be the result of an underlying R gene (Hauge et al., 2001), but was later determined to be the result of single nucleotide polymorphisms (SNPs) near putative ligand-binding sites in a serine hydroxymethyltransferase (SHMT) gene (Liu et al., 2012). SHMT is a highly conserved enzyme present across kingdoms and is responsible for the reversible conversion of serine to glycine and tetrahydrofolate (a folic acid derivative) to 5,10- methylenetetrahydrofolate. Hypotheses surrounding SHMT’s function in an incompatible response were formulated based on its crucial role in folate one-carbon metabolism and thus diminishing the syncytium’s nutrient provisions to the feeding nematode (Liu et al., 2012). It has also been noted that a disturbance in folate homeostasis leads to apoptosis of mammalian cells (Hoeferlin et al., 2013; Novakovic et al., 2006) and the altered regulation of SHMT therefore may also result in the degeneration of a syncytium (Liu et al., 2012). Alternative hypotheses include an as yet undefined regulation of defense- related genes, as the upregulation of a number of stress and defense-related genes has been observed in degenerating syncytia during an incompatible reaction (Kandoth et al., 2011). Genetic diversity between populations of SCN, as measured by their ability to grow on a set of differential lines, was observed not long after the first report of SCN in the U.S. in North Carolina in 1955 (Winstead et al., 1955). Resistance sources in available soybean cultivars was quickly identified (Ross and Brim, 1957) as was diversity in SCN populations’ ability to grow on these different resistant lines. PI 88788, for instance, was found to be resistant to an SCN isolate from Tennessee but susceptible to a North Carolina isolate (Ross, 1962, as cited in Riggs and Schmitt, 1988). Our knowledge of the diversity between populations of SCN continued to expand (Anand et al., 1994; Chen et al., 2001; Dong et al., 2005; Niblack et al.,1993; Niblack et al., 2003) as well as the genetic diversity present within populations (Colgrove et al., 2002; Dong et al., 2005; Zhang et al., 1998)

19 In work leading up to the turn of the century, SCN were classified into races according to their ability to reproduce on a chosen set of differential lines. However, populations were being tested that did not fit within the race classification system, and even as differentials and additional races were added (Riggs and Schmitt, 1988), the race designation soon became inadequate in describing or predicting a population’s virulence to any given source of resistance (reviewed in Niblack, 2004). In order to better understand the diversity in SCN populations, and the corresponding effectiveness of the sources of resistance available, an HG Type scheme (HG: H. glycines) was devised. The HG Type test, formulated by Niblack et al. (2002), describes populations of SCN in terms of their compatibility on a set of seven indicator lines, which contain the sources of resistance that have been deployed to date. Determining the HG Type of SCN allows scientists to monitor changes in populations and differences between populations, work on developing new resistant cultivars, and direct improved management strategies regarding crop rotation, specifically resistance rotation (Niblack et al., 2002). The HG Type of a given population of SCN is determined by the Female Index (FI: (average number of females on indicator line ÷ average number of females on susceptible check) ×100). A population is designated as HG Type_ based on whether FI ≥ 10 on each of the seven indicator lines (Niblack et al., 2002). For example, a population whose FI ≥ 10 only on PI 88788 (indicator line number 2), PI 209332 (indicator line number 5), and PI 548316 (“Cloud,” indicator line number 7), is designated as HG Type 2.5.7, whereas a population that does not produce a FI ≥ 10 on any of the indicator lines is designated as HG Type 0. The PI sources of resistance fall into two main groups based on their effect on SCN parasitism: PI 88788 resistance together with PI 209332 and PI 548316, and PI 548402 (Peking) resistance together with PI 90763 and PI 89772, and to some extent PI 437654 (Colgrove and Niblack, 2008). In resistance conferred by cultivar Forrest (resistance from Peking), cells surrounding syncytium die, essentially cutting them off, leading to condensed cytoplasm in and death of syncytium, whereas in cultivar Bedford (resistance from PI 88788) nuclear degeneration precedes condensing cytoplasm (Kim et al., 1987). Differences have also been noted in the nematode stage most affected; PI 209332 resistance predominantly affects the development of third and fourth stage

20 juveniles whereas Pickett (Peking) resistance affects mainly second and third stage juveniles, in addition to male development as does PI 89772 (Halbrendt et al., 1992). These groupings have also been found to correlate with the status of the Rhg1 locus, copy number and α-SNAP amino acid polymorphism (Cook et al., 2014). Interestingly, Rhg1 in PI 437654 is similar to that of PI 548402 (Peking), PI 90763, and PI 89772 in sequence and copy number. Yet Rhg1 still contributes in large part to PI 437654 resistance to an SCN population (otherwise virulent to PI 88788 Rhg1). This effect also has a significant interaction with Rhg4 (Brucker et al., 2005), an interaction that has also been observed in Peking (Meksem et al., 2001). How populations of SCN are overcoming host resistance is an important area of research, particularly due to the prevalence of virulence to the widely planted sources of resistance, such as the current over-dependence on PI 88788 in the Midwest (Colgrove and Niblack, 2008), where over 85% of SCN-resistant cultivars are derived from this line (Skorupska et al., 1994). Major genes have been discovered which are associated with the ability of SCN to grow on PI 88788, Peking, and PI 90763 (Dong et al., 1997). A study by Dong et al. (1997) created eight inbred lines of SCN, which aided in the identification of a dominant Ror locus (Ror: reproduction on a resistant host) associated with the capability of reproducing on PI 88788. PI 437654 remains the most effective and still underused source of resistance, however strains of SCN virulent to this source of resistance have also been found (Young, 1998; Niblack et al., 2003). These findings highlight the need to expand sources of resistance and manage nematode populations to prolong the effectiveness of resistance sources currently in use. While the rotation of resistance sources can be used to manage SCN infestations, it is worth noting that this does not necessarily confer stabilizing selection to slow the formation of resistance- breaking strains (Young and Hartwig, 1988). Another objective of this research was to determine potential virulence factors that have allowed SCN to successfully infect PI 88788 by examining the transcriptional profile of different HG Type populations.

21

1.7 HOST RESPONSE TO RKN & SCN PARASITISM

Successful parasitism is dependent upon more than the evasion of host defenses; RKN must also manipulate and ultimately redifferentiate root cells in order to glean the nutrition required for reproduction. One of the first signs of giant cell induction is the development of a binuclear vascular cell (de Almeida Engler, 1999), followed by additional synchronous nuclear divisions (Jones and Payne, 1978). Cytoskeleton rearrangement is important for the feeding site to develop (also required for syncytial development) as treatment with a chemical inhibitor was shown to disrupt this process and impede nematode development (de Almeida Engler, 2004). As molecular biology techniques have advanced from promoter trapping studies to whole transcriptome sequencing, progress has been made in understanding regarding the transcriptional response associated with feeding site initiation and maintenance. Jammes et al. (2005) observed that 70% of nematode-regulated defense genes were locally repressed. Other gene categories of interest include: those involved in cell wall modification such as expansins and pectinases (reviewed in Gheysen and Mitchum, 2009); in metabolism, such as RPE (an enzyme in the pentose phosphate pathway), which was determined to be essential for giant cell formation (Favery, 1998); and genes involved in cell cycle activation such as cyclins and cyclin-dependent kinases, which were differentially expressed and essential for proper feeding site establishment (de Almeida Engler et al., 1999). The RKN feeding site is thought to be symplastically isolated from the root vasculature, and it is therefore expected that the feeding site would induce the production of associated transporters necessary for it to become a nutrient sink. Transcriptional studies examining expression changes in RKN-infected versus noninfected Arabidopsis roots have found several transporters to be differentially regulated, amongst which are members of amino acid transporter families (Hammes et al., 2005; Hammes et al., 2006;

22 Jammes et al., 2005). Disruption of amino acid transport with knockout mutants of the Arabidopsis amino acid permeases AAP3 and AAP6 was found to adversely affect nematode development and result in progeny with less lipid reserves (Marella et al., 2012). Initial studies investigating the host response to SCN infection also examined transcriptional changes in whole infected root systems (Alkharouf et al., 2006; Khaan et al., 2004). Much like RKN, an important component of the host response is specific to the nematode feeding site. Other groups have investigated host responses specific to syncytia, also using laser-capture microdissection (LCM) to examine this tissue of interest (Ithal et al., 2007; Klink et al., 2010). For example, during an incompatible interaction (an avirulent SCN population on a soybean containing Rhg1), ~16% of differentially regulated genes are related to stress or defense responses; upregulated genes included those involved in the hypersensitive response and salicylic acid signaling (Kandoth et al., 2011). Ithal et al. (2007) observed a local downregulation of jasmonic acid biosynthesis, and response-related gene expression, which also indicates a local repression of defense responses. As the importance of phytohormones in normal plant growth and development has long been intensely studied, it was generally thought they would also play an important role in feeding site development. Evidence of their involvement in the formation of the feeding sites was indicated as early as the 1960’s. Balasubramanian and Rangaswami (1962) showed the presence of indole compounds (precursors of auxins) in root tissue infected with M. javanica in comparison with noninfected tissue. Since this initial discovery there have been numerous studies on the role of auxin in RKN parasitism (reviewed in Goverse and Bird, 2011). Using the auxin-responsive promoter element DR5, Karczmarek et al. (2004) discovered the feeding site-specific increase in perceived auxin concentration during early feeding site development by both M. incognita and the beet cyst nematode, BCN (Heterodera schachtii Schmidt). This was later determined to be due to the accumulation of auxin rather than to increased sensitivity to auxin signaling (reviewed in Goverse and Bird, 2011).

23 As with giant cells, the cyst-induced feeding site was also once thought to be symplastically isolated from the root vasculature. This conclusion was drawn from observations of cell wall deposits near plasmodesmata and the inability of microinjected dyes to move into cells neighboring the syncytium (Böckenhoff and Grundler, 1994). It is now understood that nutrient transport into the syncytium does in fact have a symplastic route, via plasmodesmata from the surrounding phloem (Hoth et al., 2005). The upregulation of many transporters was also observed during both BCN and SCN parasitism (Hofmann et al., 2007; Klink et al., 2005), indicating that the apoplastic route is also important to this interaction.

1.8. ROLE OF RNA SILENCING IN PLANT-PARASITIC NEMATODE RESEARCH

The effects of RNA-induced gene silencing were first discovered in the 1990’s, when research groups were attempting to overexpress a gene that would augment a flower’s pigmentation (Napoli et al., 1990; van der Krol et al., 1990). However, instead of enhanced pigmentation they observed a dramatic reduction in the expression of both the introduced and endogenous gene. This phenomenon, which they termed “co- suppression” (now referred to as RNA silencing) had important implications for future work, due to the mechanism’s ability to act in trans (Napoli et al., 1990), and at the transcriptional and post-transcriptional levels (van der Krol et al., 1990). The ability to use RNA to disrupt the expression of specific genes was first developed and utilized in the nematode (Maupas) Dougherty with the aim of creating targeted knockdowns for genetic studies (Fire et al. 1991). At the time, it was believed that the resultant silencing was chiefly due to hybridization between the introduced antisense RNA with the target gene’s mRNA; however, upon comparing the effectiveness of introducing single-stranded versus double-stranded RNA (ssRNA versus dsRNA), Fire et al. (1998) discovered that the dsRNA was a far more effective silencing trigger than both sense and antisense ssRNA. It was also observed that the reductions in transcript levels required only a few injected molecules of dsRNA, implying that a catalytic process

24 and amplification were taking place as well (Fire et al., 1998; Sijen et al., 2001). This work prompted ongoing investigations into the mechanisms and pathways responsible for the newly discovered phenomenon with new implications for how genes are regulated. One of the primary functions of RNA silencing is to defend host cells against the invading nucleic acids of viruses. In fact, multiple research groups had predicted its direct involvement in antiviral defense in the 1990s (Lindbo et al., 1993; Ratcliff et al., 1997; Smith et al., 1994). Genetic data has revealed that when certain silencing machinery components are compromised in a plant, the corresponding mutant plant becomes more susceptible to viral infection. For instance, Arabidopsis mutants, such as dicer-like 2 (dcl2) and HUA enhancer 1 (hen1) have been found to be hypersusceptible to various RNA viruses (Garcia-Ruiz et al., 2010; Zhang et al., 2012). In response, viruses have evolved to inhibit or evade this defense through the production of proteins that suppress these pathways. For example, potyviral helper-component proteinase (HC-Pro) was discovered to be a pathogenicity determinant that has a synergistic effect during co- infections with heterologous viruses through its ability to counteract RNA silencing (Kasschau et al., 2003; Pruss et al., 1997) by interfering with both short interfering-RNA (siRNA) and micro-RNA (miRNA) pathways (Ebhardt et al., 2005; Kasschau et al., 2003). Suppression of host RNA silencing, through a variety of mechanisms, is likely required for many successful viral infections (reviewed in Burgyán and Havelda, 2011; Vance and Vaucheret, 2001). More recently, non-viral pathogens have also been documented to produce effectors that suppress their host’s silencing pathways, and the roles of posttranscriptional RNA silencing in plant pathogen-host interactions have recently been reviewed (Katiyar- Agarwal and Jin, 2010; Peláez and Sanchez, 2013; Weiberg et al., 2014). Clearly, plant viruses are not alone in their production of silencing suppressors, as bacteria (Navarro et al., 2008; Wang et al., 2011) and an oomycete (Qiao et al., 2013) have also been found to utilize this virulence strategy. A well-characterized non-viral example of effector proteins that suppress host miRNA pathways comes from studies with Pseudomonas syringae van Hall strain DC3000 (Navarro et al., 2008). These effectors were found to reduce mature miRNA abundances normally activated by basal defense pathways. Concurrent work

25 with Arabidopsis mutants deficient in RNA silencing components responsible for processing miRNAs were found to be more susceptible to infection by P. syringae as well as the growth of nonpathogenic bacteria, further implicating the role of miRNAs in basal defense. The first description of a eukaryotic plant pathogen that generates silencing suppressors was Phytophthora sojae, an oomycete pathogen of soybean. Two of its effectors were shown to suppress distinct RNA silencing pathways in the host and the importance of these effects on P. sojae virulence was also illustrated (Qiao et al., 2013). One P. sojae effector induces a broad reduction in siRNA abundances while the second affects specific trans-acting siRNAs (derived from noncoding TAS genes, targeted by miRNAs). It is now anticipated that counteracting RNA silencing-mediated defense is a common strategy used by diverse pathogens to sustain their survival and reproduction. To date, there have been indications that plant-parasitic nematodes may influence silencing pathways in their host. The abundances of both siRNAs and miRNAs in Arabidopsis, as well as some of the target genes they regulate, are altered during beet cyst nematode (BCN) infection (Hewezi et al., 2008). Some of these alterations play a critical role in the developmental progression of the nematode feeding site (Hewezi et al., 2012). Altered miRNA abundances have also been observed during soybean cyst nematode (SCN) infection (Li et al., 2012). However, in both of these studies whole infected roots where analyzed, with little data suggesting that the alteration of RNA silencing pathways was specific to the nematode feeding site. Biotrophic pathogens, such as RKN, are of particular interest because unlike necrotrophic pathogens, they require living host tissue to complete their life cycle. After penetrating a root, RKN initiates the formation of a feeding site of about 5-7 giant cells. RKN essentially reprogram these host cells into nutrient sinks to feed from, allowing the nematodes to be sedentary for the remainder of their lifecycle. The success of a pathogen- host interaction depends on a pathogen’s ability to obtain nutrients from its host while avoiding host defenses. Biotrophic pathogens require strategies to obtain nutrition from their host without becoming targeted by these defense responses. An objective of this work was to determine whether RKN interfere with their host’s RNA silencing

26 pathways, and whether this interference benefits the infection. Using RNA silencing pathways has also become a useful tool for the functional analysis of genes involved in parasitism in plant-parasitic nematodes. This method has been successfully used in the creation of thousands of gene knockdowns in the free-living nematode model system Caenorhabditis elegans (Fraser et al., 2000; Gonczy et al., 2000; Maeda et al., 2001) targeting nearly every gene in the genome (Kamath and Ahringer, 2003). The obligate biotrophy of both RKN and SCN present an added challenge to functional genetic studies. One approach involves inducing the uptake of dsRNA by infective J2 by soaking them in a solution with compounds such as resorcinol or octopamine (Rosso, 2005; Urwin et al., 2002). This method has been shown to successfully knockdown transcripts of interest; however, the effectiveness of this approach has been somewhat limited to early stages of infection. In order to determine the function of genes involved in parasitism after the initial stages of infection, researchers have begun to express dsRNA and miRNA in planta, targeting nematode transcripts of interest. A compilation of the results from such studies is provided in Table 1.1. Aside from functional analyses, this technique has attractive potential as a new method for nematode control. Our studies and results discussed in Chapter 2 (Root-Knot Nematode Parasitism Suppresses RNA Silencing) indicate that the effectiveness of these methods may be improved with more research on effective delivery strategies.

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36 Table 1.1. Summary of method, target, and results of studies utilizing RNAi for plant-parasitic nematode control.

Nematode Host Delivery Target RNA Validation Effect on Infection Reference unc-15 (paramyosin) dsRNA / siRNA in planta dec. lesion area D. destructor sweet potato dsRNA - transgenic Fan et al. 2015 (C. elegans lethal) mRNA dec. in nematode inc. yield in infected field Hg-rps-23 H. glycines soybean dsRNA soak/uptake by J2 mRNA dec. in nematode loss of movement in J2 Alkharouf et al. 2007 (C. elegans lethal) dec. mature females and H. glycines soybean dsRNA soak/uptake by J2 hg-syv46 mRNA dec. in nematode Bakhetia et al. 2007 dec. proportion females:males initial inc. in mRNA dg13 (secreted from dorsal abundance observed for dec. female development and H. glycines soybean dsRNA soak/uptake by J2 Bakhetia et al. 2008 gland) some; mRNA dec. in dec. proportion females:males nematode H. glycines soybean dsRNA - transgenic Major sperm protein ACT2 not detected dec. number of eggs Steeves et al., 2006 H. glycines soybean Ribosomal protein 3a,4. dsRNA – transgenic roots not detected dec. number of females Klink et al., 2009 Spliceosomal SR protein H. glycines soybean dsRNA – transgenic roots Fib1,Y25 mRNA dec. in nematode dec. number of eggs Li et al., 2010 H. glycines soybean dec. J2 invasion (30-39%) dsRNA – transgenic roots Pectate lyase: HG-PEL-6 mRNA dec. in nematode dec. female development (36- Peng et al. 2016 45%) H. glycines parasitism genes: H. schachtii Arabidopsis dsRNA - transgenic mRNA dec. in nematode dec. mature females (23-64%) Sindhu et al. 2008 3B05, 4G06, 8H07, 1006 Arabidopsis Mc16D10L (secreted dec. egg masses (57-67%) M. chitwoodi dsRNA - transgenic mRNA dec. in nematode Dinh et al. 2014 potato parasitism gene) and in potato (63-71%) 37 M. incognita dec. J2 infectivity M. javanica dsRNA soak/uptake by J2 16D10 (secreted parasitism dsRNA / siRNA in planta Arabidopsis dec. number of galls (63-90%) Huang et al. 2006 M. arenaria dsRNA - transgenic gene) mRNA dec. in nematode dec. egg production (69-93%) M. hapla Miduox (dual oxidase) Arabidopsis dsRNA soak/uptake by J2 dec. J2 retention M. incognita Mispc3 (signal peptidase dsRNA in planta Charlton et al. 2010 red bean dsRNA - transgenic dec. mature females (52-63%) complex 3) Tyrosine Phosphate, dec. number of galls M. incognita soybean dsRNA – transgenic roots itochondrial stress -70 protein mRNA dec. in nematode Mathews et al., 2010 dec. female diamter precursors MSP M. incognita tomato dsRNA – transgenic roots Rpn7 mRNA dec. in nematode dec. number of eggs Niu et al., 2012 M. incognita dsRNA soak/uptake by J2 peroxiredoxin – Mi-prx2.1 not detected dec. number of galls (60%) Dubreuil et al., 2011 MjTis11 (putative siRNA in planta nsd. on fecundity or egg hatching M. javanica tobacco dsRNA - transgenic Fairbairn et al. 2007 transcription factor) mRNA dec. in nematode rate M. artiellia wheat dsRNA soak of egg mass (chitin synthase) mRNA in egg masses delayed hatching Fanelli et al. 2005 Continued 37 Table 1.1. continued

Nematode Host Delivery Target RNA Validation Effect on Infection Reference TP (tyrosine phosphatase) mRNA dec. in infected dec. number of galls (92/94%) M. incognita soybean dsRNA - transgenic MSP (mitochondrial stress-70 Ibrahim et al. 2011 tissue dec. female size precursor) M. incognita dec. number of galls Arabidopsis dsRNA - transgenic Parasitism gene 8D05 not detected [Xue et al.,2013]

M. incognita dec. number of galls Arabidopsis dsRNA - transgenic Calreticulin - MiCRT dec. mRNA in nematode [Jaouannet et al. 2013]

Cpn-1, Y25, Prp-17 siRNA in planta dec. egg production (79-95%) H. glycines soybean dsRNA - transgenic (related to reproduction or mRNA dec. in nematode for [Li et al. 2010] dec. number of cysts fitness) later two genes

cysteine proteinase dec. female:male ratio H. glycines soybean dsRNA soak/uptake by J2 dec. mRNA in nematode [Urwin et al. 2002] G. pallida potato C-type lectin dec. number of cysts major sperm protein (MSP) nsd. splicing factor protein dec. galls M. incognita tobacco dsRNA - transgenic dec. mRNA in nematode [Yadav et al. 2006] integrase dec. egg production

38

38

CHAPTER 2: ROOT-KNOT NEMATODE PARASITISM SUPPRESSES RNA SILENCING

2.1. ABSTRACT

Root-knot nematodes damage crops around the world by developing complex feeding sites from normal root cells of their hosts. The ability to initiate and maintain this feeding site (composed of individual “giant cells”) is essential to their parasitism process. RNA silencing pathways in plants serve a diverse set of functions, from directing growth and development to defending against invading pathogens. Influencing a host’s RNA silencing pathways as a pathogenicity strategy has been well documented for viral plant pathogens, but recently it has become clear that silencing pathways also play an important role in other plant pathosystems. To determine if RNA silencing pathways play a role in nematode parasitism, we tested the susceptibility of plants that express a viral suppressor of RNA silencing. We observed an increase in susceptibility to nematode parasitism in plants expressing viral suppressors of RNA silencing. Results from studies utilizing a silenced reporter gene suggest that active suppression of RNA silencing pathways may be occurring during nematode parasitism. With these studies we are the first to demonstrate the suppression of RNA silencing during the interaction between a plant-parasitic animal and its host.

39

2.2. INTRODUCTION

Root-knot nematodes (RKN), belonging to the genus Meloidogyne, represent one of the most economically damaging genera of plant-parasitic nematodes worldwide (Sasser, 1980). RKN parasitism is dependent upon the nematode’s ability to initiate and maintain its feeding site (composed of individual “giant cells”) (Bird, 1962). In order to manipulate the normal physiology of those host cells, RKNs secrete effector proteins that influence several pathways including those related to cell cycle, metabolism, cell wall modification, and defense (Hewezi and Baum, 2013). RNA silencing pathways endogenous to plants have an important regulatory role in many of these pathways (reviewed in Baulcombe, 2004; Chen, 2009; Jones-Rhoades et al., 2006; Vance and Vaucheret, 2001). Influencing a host’s RNA silencing pathways as a pathogenicity strategy has been well documented for viral plant pathogens (Chellappan et al., 2004; Li et al., 1999; Ratcliff et al., 1997; Vance and Vaucheret, 2001), but recently it has become clear that silencing pathways also play an important role in other plant pathosystems (Qiao et al., 2013; Weiberg et al., 2013). RNA silencing monitors the intracellular occurrence of double-stranded RNA (dsRNA), which can be of endogenous or exogenous origins, and processes dsRNAs into small RNAs of discrete sizes (21- 25 nucleotides [nt]) termed small interfering RNAs (siRNAs). siRNAs then mediate the silencing of homologous genetic elements by guiding RNA-induced silencing complexes (RISCs) to complementary RNA or DNA, and modulate the expression of the corresponding gene products. One of the primary functions of RNA silencing is to defend host cells against the invading nucleic acids of a virus. In fact, multiple research groups had predicted its direct involvement in antiviral defense in the 1990s (Lindbo et al., 1993; Ratcliff et al., 1997; Smith et al. 1994). Genetic data has revealed that when certain silencing machinery components are compromised in a plant, the corresponding mutant plant becomes more susceptible to viral infection. For instance, Arabidopsis mutants, such as dicer-like 2 (dcl2) and HUA enhancer 1 (hen1)

40 have been found to be hypersusceptible to various RNA viruses (Garcia-Ruiz et al. 2010; Zhang et al. 2012). In response, viruses have evolved to inhibit or evade this defense through the production of proteins that suppress these pathways. For example, potyviral helper-component proteinase (HC-Pro) was discovered to be a pathogenicity determinant that causes a synergistic effect during co-infections with heterologous viruses through its ability to counteract RNA silencing (Kasschau et al., 2003; Pruss et al., 1997), interfering with both short interfering-RNA (siRNA) and micro-RNA (miRNA) pathways (Ebhardt et al., 2005; Kasschau et al., 2003). It has since been concluded that suppression of host RNA silencing, through a variety of mechanisms, is likely required for many successful viral infections (reviewed in Burgyán and Havelda, 2011; Vance and Vaucheret, 2001). More recently, non-viral pathogens have also been documented to produce effectors that suppress their host’s silencing pathways, and the roles of posttranscriptional RNA silencing in plant pathogen-host interactions have recently been reviewed (Katiyar- Agarwal and Jin, 2010; Peláez and Sanchez, 2013; Weiberg et al., 2014). It has become clear that plant viruses are not alone in their production of silencing suppressors, as bacteria (Navarro et al., 2008; Wang et al., 2011) and an oomycete (Qiao et al., 2013) have also been found to utilize this virulence strategy. A well-characterized non-viral example of an effector protein that suppresses host miRNA pathways comes from a study with Pseudomonas syringae strain DC3000 (Navarro et al., 2008). These effectors were found to reduce mature miRNA abundances normally activated by basal defense pathways. Concurrent work with Arabidopsis mutants, deficient in RNA silencing machinery components responsible for processing miRNAs, found them to be more susceptible to infection by P. syringae, further implicating the role of miRNAs in basal defense. The first description of a eukaryotic plant pathogen that generates silencing suppressors was Phytophthora sojae, a pathogen of soybean. Two of its effectors were shown to suppress distinct RNA silencing pathways in the host and the importance of these effects on P. sojae virulence was also illustrated (Qiao et al., 2013). One P. sojae effector induces a broad reduction in siRNA abundance while the second affects specific trans-acting siRNAs (derived from noncoding TAS genes, targeted by miRNAs). It is now

41 anticipated that counteracting RNA silencing-mediated defense is a common strategy used by diverse pathogens to sustain their survival and reproduction. To date, there have been indications that plant-parasitic nematodes may influence silencing pathways in their host. The abundances of both siRNAs and miRNAs in Arabidopsis, as well as some of the target genes they regulate, are altered during beet cyst nematode (BCN, Heterodera schachtii) infection (Hewezi et al., 2008). Some of these alterations play a critical role in the developmental progression of the nematode feeding site (Hewezi et al., 2012). Altered miRNA abundance has also been observed during soybean cyst nematode (SCN) infection (Li et al., 2012). However, in both of these studies whole infected roots where analyzed, with little data suggesting that the alteration of RNA silencing pathways was specific to the nematode feeding site. The objectives of this study were to determine whether hosts impaired in their RNA silencing pathways are hypersusceptible to nematode infection and whether M. incognita parasitism interferes with these pathways.

2.3. MATERIALS & METHODS

2.3.A. Susceptibility bioassays. Wild type (Nicotiana tabacum cv. Xanthi) and transgenic tobacco expressing Tobacco etch virus’ P1/HC-Pro (line X-27-8, created by Mallory 2001) were grown in cones with Turface-sand (1:1 v/v) (Turface MVP; Buffalo Grove, IL) amended with slow-release fertilizer. Nicotiana benthamiana lines expressing PVA HC-Pro (Lakatoset et al., 2004), CRV P19 (Savenkov and Valkonen, 2001) and TCV CP (F. Qu, personal communication) were treated similarly. M. incognita population was maintained on tomato (Solanum lycopersicum, cv. Moneymaker). Plants were each infested with 1,000 M. incognita eggs/cone. After 5 weeks, roots were cut at the crown, rinsed with water and fresh root weight was recorded. Roots were cut into approximately one inch pieces and submerged in 5% bleach for 15 minutes while gently agitated. Three 15μl subsamples were counted for each plant. Three independent experiments were completed with TEV

42 HC-Pro each including ≥ 19 plants per line. For N. benthamiana ≥ 24 plants per line were analyzed; three independent experiments were completed with TCV CP and CRV P19 and two independent experiments were completed with PVA HC-Pro.

2.3.B. Transgenic Lines. Tobacco plants (Nicotiana tabacum cv. Xanthi) homozygous for a single-locus T- DNA insertion line containing the GUS/dsRNA-silenced GUS (Fig. S3A) construct were used in this experiment. The tobacco line (Nicotiana tabacum cv. Xanthi) expressing miRNA-silenced GUS was generated as the progeny of two homozygous tobacco lines, one with a single-locus T-DNA insertion expressing the miRNA and the other a single- locus T-DNA insertion expressing GUS (Fig. S3C). The miRNA was designed using miRNA156 as a template, replacing the 22 nt binding region with sequence specific for the GUS transcript.

2.3.C. GUS-restoration pictures. Root tissue was harvested (4, 7, 14, and 21 days post infestation with juvenile RKN) and placed into ½ strength X-Gluc (Gold Biotechnology, St. Louis, MO) staining solution (Jefferson 1987) for 15 hours at 37°C. Knots were harvested and put in a fixative solution (3% v/v glutaraldehyde, 0.03% Triton X-100, 100 mM Tris pH 7.0) overnight at 4°C (Hammes et al. 2006). Knots were then placed into 3% agarose and allowed to solidify at 4°C. Approximately 80μm sections were cut using an EMS-4000 Automatic Oscillating Tissue Slicer, put onto slides and visualized with an inverted Leica DM IRB microscope equipped with a Q Imaging Retiga 2000 cooled digital camera.

2.3.D. GUS-restoration RT-PCR. Additional fresh tissue (RKN-infected and non-infested control) was simultaneously harvested and immediately frozen in liquid N2. Three root samples (each from three pooled plants) were harvested for knot tissue and uninfected tissue (surrounding knots), and non-infected tissue (tissue excluding root tips) at 7, 14, and 21dpi with J2 RKN. Total RNA was extracted with TRIZOL (Life Technologies,

43 Carlsbad, CA) and treated with RQ1 DNase (Promega, Madison, WI); measurements of quantity and quality were obtained with a Qubit® 2.0 Fluorometer (Life Technologies, Carlsbad, CA) and a Bioanalyzer (Agilent Technologies, Palo Alto, CA) respectively. RNA template (500ng) was used for cDNA synthesis following the manufacturer’s instructions with the GoScriptTM Reverse Transcription System (Promega, Madison, WI)

primed with oligo(dT)15. During cDNA synthesis a no-RT reaction was included for each RNA sample in order to detect contaminating genomic DNA. cDNA template was diluted 1:2 and amplified using Phusion® High Fidelity DNA Polymerase (New England Biolabs, Ipswitch, MA) with GUS-specific primers 5’-GCCAGTGGCGCGA- AATATTCCCGTG (sense) and 5’- GTGCTGCGTTTCGATGCGGTCACTC (antisense) with the following cycle conditions: initial denaturation at 98°C for 30s, 30 cycles of 98°C for 10s, 73°C for 20s, 72°C for 15s, with a final extension at 72°C for 10min. Protein phosphatase 2A (PP2A; 123bp region) was chosen as a gene of reference due to its relative stability shown in work with viral infections in Nicotiana benthamiana (Liu et al., 2012).

2.3.E. Statistical analysis. Egg count data from susceptibility bioassays were analyzed with JMP software (version 11.0.0; SAS Institute Inc., Cary, NC). Median egg counts of wild type and TEV HC-Pro-expressing N. tabacum were compared with the nonparametric test, Wilcoxon rank-sum (p < 0.01). Differences between means of wild type and VSR-expressing N. benthamiana were analyzed with the nonparametric test for multiple comparisons using Steel's Multiple Comparison Wilcoxon Tests (comparisons of HC-Pro and CP with the control, p < 0.025).

44 2.4. RESULTS

2.4.A. Susceptibility of plants expressing a VSR. Viral suppressors of RNA silencing (VSR) have been found to act as important virulence factors in multiple plant pathosystems. In previous studies, VSRs have been used to determine whether silencing suppression affects a pathogen’s virulence. Using a transgenic tobacco expressing the known viral suppressor P1/HC-Pro (from Tobacco etch virus (TEV), created by Mallory et al., 2001), we examined whether transgenic expression of this viral protein also rendered plants more susceptible to root-knot nematode parasitism. As shown in Fig. 1, more eggs were produced on the HC-Pro- expressing tobacco in comparison with the wild type control. Concerned that HC-Pro- expressing tobacco’s smaller root system and developmental delays could be confounding these results, we included the progeny of a dsRNA-silenced GUS x HC-Pro cross (Fig. 2.2), as their phenotype is closer to that of wild type plants. These plants, hemizygous for HC-Pro, also proved to be hypersusceptible to M. incognita infection (Fig. 2.3). We similarly tested the susceptibility of transgenic Nicotiana benthamiana lines expressing Potato virus A (PVA) HC-Pro (Savenkov and Valkonen, 2001), Turnip crinkle virus (TCV) CP (F. Qu, personal communication), or Cymbidium ringspot virus (CRV) P19 (Lakatos et al., 2004) and also observed an increase in susceptibility in the presence of two of these VSRs (Fig. 2.4); however, transgenic plants expressing P19 did not show a significant change in susceptibility. These results indicate that a host compromised in its silencing pathways is likely to be more susceptible to M. incognita infection.

2.4.B. Reversal of silencing assay. In order to address the question of whether M. incognita interfere with their host’s RNA silencing pathways, we adapted a method widely used by plant virologists, a reversal of silencing assay (Anandalakshmi et al., 1998). We developed a transgenic tobacco line with a single T-DNA insertion containing both the reporter enzyme β- glucuronidase (GUS) and a dsRNA-generating sequence corresponding to a portion of

45 the GUS transcript (Fig. 2.5). Qiao et al. (2013) utilized a similar method to illustrate P. sojae effectors’ interference of RNA silencing, indicated by restoration of a reporter gene’s expression. The figwort mosaic virus (FMV) promoter was chosen to drive the expression of both dsRNA-GUS and full length GUS due to its strong expression in roots (Govindarajulu et al., 2008) and during M. incognita infestation (Collier et al., 2005). The root system of the dsRNA-silenced GUS tobacco line created was void of any evident GUS expression (Fig. 2.6A). To verify that GUS was indeed being expressed and post-transcriptionally silenced, the line was crossed with the transgenic tobacco expressing the known viral suppressor HC-Pro (Mallory et al., 2001). Several groups have utilized this strategy to illustrate HC-Pro’s ability to suppress the silencing of a transgene (Anandalakshmi et al., 1998; Kasschau and Carrington, 1998; Mallory et al., 2001). As expected, progeny displayed a complete recovery of GUS expression (Fig. 2.6B). Tobacco plants expressing dsRNA-silenced GUS were infested with second stage juvenile (J2) root-knot nematodes; resulting root knots were stained (Fig. 2.6) and sectioned over the course of infection. From these sections it was evident that silencing of the GUS transcript was being suppressed specifically within the nematodes’ giant cells (Fig. 2.7). GUS expression was restored in the nematode feeding site as early as four days post infestation (dpi) and continued for at least 21 dpi. Reverse transcriptase PCR (RT- PCR) was performed on RNA collected from infected root tissue using primers specific to the 5’ and 3’ end of the GUS gene. RT-PCR confirmed the presence of intact GUS transcript in nematode-infected tissue (Fig. 2.8) during nematode parasitism (data shown from 7 dpi, similar results were observed at 14 and 21 dpi, Fig. 2.9). To ensure that the change in RNA silencing was not due to aberration or alterations using the same promoter sequences to drive GUS and the dsRNA-GUS constructs, we developed and tested tobacco lines that drove GUS expression using a pine Super Ubiquitin (SU) promoter (Fig. 2.5B). Like the FMV-driven GUS construct, GUS expression driven by the SU promoter occurred throughout the entire root and during nematode infection (data not shown). Progeny of the cross between the dsRNA-GUS and SU-GUS transgenic tobacco lines showed no GUS expression in the root; however, when

46 the roots were infested with RKN, GUS expression was restored in the nematode feeding site (Fig. 2.10A). Our previous test showed that the dsRNA-directed (siRNA) silencing pathways appear to be altered during nematode parasitism. We wanted to determine if the miRNA pathways might be altered as well. A miRNA-GUS construct (Fig. 2.5C) was transformed into tobacco, these lines were crossed with the pSU-GUS transgenic plant, and the progeny were tested for GUS expression. In non-infected plants, GUS expression was silenced throughout the entire root system. When M. incognita were added, GUS expression was observed in the nematode feeding site (Fig. 2.10B). This suggests that mechanisms regulating both dsRNA and miRNA pathways are altered during M. incognita infection and that these alterations are specific to the nematode feeding site.

2.5. DISCUSSION

A diverse range of mechanisms has so far been characterized for VSRs (Díaz- Pendón and Ding, 2008). Among the VSRs tested in this study, HC-Pro (TEV; N. tabacum), HC-Pro (PVA; N. benthamiana) and CP (TCV; N. benthamiana), increase host susceptibility to RKN. While HC-Pro and CP have both been shown to suppress silencing pathways, unlike HC-Pro, CP has been found to interfere at earlier stages (dsRNA processing) (Qi et al., 2004, Qu et al., 2003; Thomas et al., 2003). Although interfering at different points in silencing pathways, these two unrelated viruses both interact with a common host transcription factor (RAV2) (Endres et al., 2010) resulting in transcriptional changes diverted from defense. Unlike HC-Pro and CP, we did not observe an alteration in M. incognita susceptibility in P19-expressing plants. Also dissimilar to the other VSRs tested, P19’s primary mode of suppressing silencing is the sequestration of siRNAs (Lakatos et al., 2004). Comparing direct mechanisms of these different VSRs may help clarify why some VSRs lead to increases in susceptibility; however, the resulting effects on downstream host pathways may prove just as insightful. For example, crosstalk between small RNA pathways and jasmonate signaling have been

47 reported (Endres et al., 2010). Results from our reversal of silencing assay additionally suggest that RKN’s suppression interferes with both si- and miRNA-directed silencing pathways. It is possible that M. incognita parasitism interferes with mechanisms or substrates common between pathways. Future work should explore the interaction with more VSRs with varying mechanisms and downstream effects. Additionally, examination of the nematode susceptibility of mutant plants that are missing individual components or entire RNA silencing pathways may yield further clues into the nature of suppression of RNA silencing during nematode parasitism. Our studies using transgenic plants expressing VSRs showed an increase in nematode susceptibility. This increase in susceptibility aligns with multiple studies done in the 1960-70s showing that some virus-infected plants coinfected with root-knot nematodes produce higher levels of nematode offspring than non-virus-infected plants (Powell 1971). Conclusions drawn from these earlier studies focused on the basic biology of these organisms. Both nematode and viral pathogens are obligate biotrophs and thus sensitive to changes in their host’s physiology. The results from our experiments would suggest that it is the virus encoded VSRs that are responsible for the increase in nematode susceptibility observed in those earlier studies. Investigating nematode susceptibility of plants infected with viruses that have mutated silencing suppressors will help decipher VSRs’ role during nematode parasitism. It is also possible that some nematodes may actually benefit from their associated viruses. In the virus-transmitting Adenophorea nematodes (Xiphinema spp., Longidorus spp., spp. and Paratrichodorus spp.) we speculate that the plant viruses, previously shown to be non-essential for nematode parasitism, may in fact alter the host plant and enhance the ability of the nematode to grow and reproduce. Future experiments could examine these relationships for any effects on nematode parasitism or feeding site productivity. Aside from diseases caused by nematode-vectored viruses, nematode- associated viruses have been largely under-explored. The recent discoveries of viral genomes in a plant-parasitic nematode (Bekal et al., 2014; Bekal et al., 2011) may provide additional opportunities to explore these relationships for positive effects on nematode fitness.

48 As evidence for further investigation, there are many studies that have examined changes in gene expression in Arabidopsis over the course of RKN and BCN infection. A tool was developed by Cabrera et al. (2014) that combines data from a few of these studies in a searchable format: NEMATIC (NEMatode-Arabidopsis Transcriptomic Interaction Compendium). The expression changes in RNAi machinery and targets during M. incognita infection are modest but significant and do display interesting trends (Table 2.1). Of the differentially expressed RNA silencing machinery genes, 7/8 are upregulated in M. incognita galls/giant cells and 13/14 are upregulated in syncytia (Jammes et al., 2005; Szakasits et al., 2009). In our own microarray dataset comparing expression in giant cells versus surrounding non-giant cells (Morse et al., 2010), we observe similar trends in expression (Table 2.1) wherein 24/25 RNA silencing machinery genes are upregulated. These latter two studies (Szakasits et al., 2009 and Morse et al., 2010) observed more differentially expressed genes in these pathways than that examining gall tissue (Jammes et al., 2005), which may reflect expression changes exclusive to syncytia and giant cells. A majority of differentially expressed ta-siRNA- targeted genes are also upregulated in the syncytium and giant cell-specific studies, providing further indication that disruptions to these pathways are specific to the nematode feeding site. Our data showing changes in nematode susceptibility in VSR plants along with the reporter gene studies showing suppression of RNA silencing in giant cells combined with the data gleaned from existing nematode-induced gene-expression studies indicates that there is more to be learned about how RNA interference is being suppressed during nematode parasitism. While this work illustrates that M. incognita parasitism suppresses host RNA silencing and that this suppression appears to be important to a successful infection, the mechanism responsible remains elusive. M. incognita are known to produce and secrete a plethora of proteins and small molecules during infection (Hewezi and Baum 2013; Hussey 1989), one of which could be directly interfering with silencing components much like those discovered in other pathosystems. However RKN’s obligatory biotrophic lifestyle may offer other mechanisms of interference, unique to their induction of physiologically altered and maintained host cells. Further elucidation of

49 pathway components with which M. incognita parasitism is interfering and how this effect is engendered will expand our understanding of the roles of RNA silencing during parasitic interactions.

2.6. ACKNOWLEDGEMENTS

We acknowledge Dr. Zhi Qi for creating the miRNA GUS construct, Kevin Lutke (Donald Danforth Plant Science Center, St. Louis, MO) for creating the transgenic tobacco plants, and Dr. Feng Qu for providing the Nicotiana benthamiana plants used in this study. This work was supported in part by funding from NSF: #IOS-1104334 and The Ohio State University, Department of Plant Pathology and Center for Applied Plant Sciences, SoyRes Research Team.

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55

160 * 140

120

(thousands) 100

80

60

40

20 RKN eggs / gram root root / gram eggs RKN

0 WT HC-Pro N. tabacum

Figure 2.1. Tobacco expressing the TEV viral suppressor, HC-Pro, are more susceptible to M. incognita (RKN) infection. Data represent the mean ± s.e.m., n ≥ 19 plants; results were repeated in 2 additional replications (Wilcoxon, * p < 0.01).

56 A B

Figure 2.2. Relative absence of β-1, 4-endoglucanase (GUS) expression in dsRNA:GUS two-week old seedlings (A) and restoration of GUS expression in progeny (B) of the dsRNA:GUS line crossed with P1/HC-Pro (line X-27-8, created by Mallory, 2001). The reciprocal cross also shows a restoration of GUS (not shown).

57 900 900 dsRNA:GUS * Wild Type 800 800 HC-Pro dsRNA:GUS x 700 700 HC-Pro * 600 600 * 500 500 * 400 400

300 * 300 200 * 200 100 100

% Eggs/g root relative control to relative root Eggs/g % 0 0 Rep 1 Rep 2 Rep 3 Rep 1 Rep 2 Rep 3 Independent Replications Independent Replications

Figure 2.3. Susceptibility of tobacco plants either homozygous or hemizygous for HC- Pro is enhanced. HC-Pro-expressing tobacco are significantly more susceptible to M. incognita than wild type in three independent experiments (data represent mean ± s.e.m., n ≥ 19, Wilcoxon * p < 0.01). Tobacco hemizygous for HC-Pro are also significantly more susceptible than the parent lacking HC-Pro (data represent mean ± s.e.m., n ≥ 15, Wilcoxon * p < 0.008).

58 50

45 * 40 * 35 30 25 20 15 10 5 RKN Eggs / gram root (thousands) root gram / RKN Eggs 0 WT HC-Pro CP P19 N. benthamiana

Figure 2.4. Two of three VSR-expressing Nicotiana benthamiana are more susceptible hosts to M. incognita. Data represent the mean ± s.e.m., n ≥ 24 plants; results were repeated in two additional replications with TCV CP and CRV P19 and one additional replication with PVA HC-Pro (Steel's Multiple Comparison Wilcoxon Tests, * p < 0.01).

59 A

B

C crossed with:

Figure 2.5. Silenced-reporter gene constructs are illustrated above. A) Expression cassette for dsRNA-silenced GUS. B) Individual expression cassettes for dsRNA- silenced GUS and SU-driven GUS. C) Individual expression cassettes for miRNA- silenced GUS and SU-driven GUS.

60 A B

Figure 2.6. Histochemical analysis of X-gluc stained dsRNA:GUS tobacco roots visualized with a dissecting microscope. Non-infected roots (A) and roots 14 dpi with J2 M. incognita wherein recovery of GUS expression is evident in knot tissue in close proximity to female M. incognita (B). Knot tissue was harvested for sectioning.

61 Figure 2.7. Restoration of dsRNA-silenced β-1, 4-endoglucanase (GUS) expression in nematode feeding sites. Sections taken at A) 4, B) 7, C) 14, and D) 21 days post inoculation with second-stage juvenile worms; GUS restoration evident in nematode (red*) giant cells (white*) (scale bar = 100 μm).

62 Figure 2.8. Restoration of β-1, 4-endoglucanase (GUS) transcripts in nematode feeding sites, 7 dpi with juvenile M. incognita. RT-PCR products produced from uninfected and infected (knot) tissue; shown are three technical replications each representing three pooled plant samples. Note absence of GUS amplicon in uninfected tissue, and presence of GUS amplicon in knot tissue. Corresponding PP2A reference gene amplicon is presented along with no template control (NTC). Similar results were observed at 14 and 21dpi and repeated in three independent experiments.

63 1 2 3 4 5 6 7 8 9 10 1 2 3 4 5 6 7 8 9 10

GUS

PP2AA

A B

GUS

PP2AA C

Figure 2.9. RT-PCR products from non-inoculated tissue (lanes 1-3), noninfected tissue (lanes 4-6), knot tissue (lanes 7-9), and no template control (lanes 9). 1 kb β-1, 4- endoglucanase amplicon present in knot tissue at 7 (A), 14 (B), and 21dpi (C). Three technical replicates shown each from three pooled plant samples. PP2A 123bp amplicon from each corresponding sample in lower gel (100bp ladder, NEB).

64 *

* *

A B

Figure 2.10. Restoration of silenced β-1, 4-endoglucanase (GUS) in nematode feeding sites. A) Early and late (inset) during M. incognita (*) infection in dsRNA-silenced SU- GUS whole roots. B) Cross-section of 21dpi M. incognita (*) feeding site in miRNA- silenced GUS roots.

65 RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT1G01040 0.93 N/A N/A N/A RNA silencing AT1G05460 0.96 0.70 0.73 N/A machinery AT1G09700 N/A N/A N/A 1.10

AT1G14790 -0.77 -1.72 -2.63 N/A

AT1G24450 N/A 0.87 1.00 N/A

AT1G24706 N/A N/A N/A N/A

AT1G30070 2.06 N/A N/A 4.60

AT1G31280 N/A N/A N/A N/A

AT1G31290 N/A N/A N/A 1.30

AT1G32490 0.47 N/A N/A N/A

AT1G33060 N/A N/A N/A N/A

AT1G48410 0.56 0.67 N/A N/A

AT1G69440 N/A N/A N/A N/A

AT1G73840 0.78 N/A N/A N/A

AT1G75660 N/A N/A N/A N/A

AT2G05370 N/A N/A N/A N/A

AT2G13540 1.04 N/A N/A N/A

AT2G16390 N/A N/A N/A 0.50

AT2G19430 N/A N/A N/A N/A

AT2G27040 0.65 0.92 1.19 1.50

AT2G27100 1.13 N/A N/A N/A

AT2G27880 1.84 N/A N/A N/A

AT2G28380 0.89 N/A N/A N/A

AT2G40030 1.42 N/A N/A N/A

AT3G03300 N/A N/A N/A N/A

AT3G05040 N/A N/A N/A N/A

Continued

66 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT3G15390 N/A N/A N/A N/A RNA silencing AT3G20420 N/A N/A N/A 1.10 machinery AT3G20550 N/A N/A N/A N/A

AT3G26932 N/A N/A N/A N/A

AT3G43920 N/A N/A 0.86 1.60

AT3G49500 0.64 0.78 1.05 N/A

AT3G62800 1.02 N/A N/A 1.46

AT4G08980 N/A 0.73 N/A N/A

AT4G11130 N/A -1.80 -1.67 N/A

AT4G20910 N/A N/A N/A 0.60

AT4G37510 N/A N/A N/A 1.20

AT5G01400 0.83 N/A N/A N/A

AT5G04290 1.15 1.32 1.55 N/A

AT5G05540 N/A 0.69 0.64 N/A

AT5G09860 0.45 N/A N/A N/A

AT5G14620 N/A N/A N/A N/A

AT5G16790 N/A N/A N/A 0.60

AT5G18830 0.44 N/A N/A -1.00

AT5G20320 N/A 0.72 0.88 N/A

AT5G21150 3.94 N/A N/A 0.40

AT5G23570 N/A N/A 0.76 1.40

AT5G42920 N/A N/A N/A N/A

AT5G43810 1.73 N/A N/A 1.90

AT5G44200 0.44 N/A N/A 2.00

AT5G49160 N/A N/A N/A N/A

AT5G56130 0.84 N/A N/A N/A

Continued 67 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT5G63110 0.55 N/A N/A N/A RNA silencing AT5G63980 N/A N/A N/A N/A machinery AT5G66750 1.38 N/A N/A N/A

AT5G67240 N/A N/A N/A N/A

AT1G01040 0.93 N/A N/A N/A miRNA targets AT1G02860 0.84 N/A N/A -1.20

AT1G06580 N/A N/A N/A N/A

AT1G12210 N/A N/A N/A N/A

AT1G12220 N/A N/A N/A N/A

AT1G12280 N/A N/A N/A 0.70

AT1G12290 N/A N/A N/A -1.30

AT1G12820 0.62 N/A N/A -0.80

AT1G15890 N/A N/A N/A -1.20

AT1G17590 N/A N/A N/A N/A

AT1G22960 1.07 N/A N/A N/A

AT1G24881 N/A N/A N/A N/A

AT1G27340 -1.24 N/A N/A N/A

AT1G27360 N/A N/A N/A N/A

AT1G27370 0.94 N/A N/A N/A

AT1G30210 1.52 N/A N/A N/A

AT1G30330 N/A N/A 1.14 0.90

AT1G30490 1.57 N/A 0.91 2.00

AT1G31280 N/A N/A N/A N/A

AT1G31840 N/A N/A N/A N/A

AT1G48410 0.56 0.67 N/A N/A

AT1G51480 N/A N/A N/A N/A

Continued 68 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT1G52050 N/A N/A N/A N/A miRNA targets AT1G52060 -2.57 N/A N/A N/A

AT1G52150 N/A N/A N/A N/A

AT1G53160 N/A N/A N/A N/A

AT1G53230 N/A -0.76 N/A N/A

AT1G53290 N/A N/A N/A 2.00

AT1G54160 N/A N/A N/A 2.70

AT1G56010 -2.71 -1.13 -0.83 -4.80

AT1G57570 N/A N/A N/A -1.20

AT1G62590 0.80 N/A N/A N/A

AT1G62630 N/A N/A N/A N/A

AT1G62670 N/A N/A N/A N/A

AT1G62720 0.76 N/A N/A N/A

AT1G62860 1.60 N/A N/A N/A

AT1G62930 N/A N/A N/A N/A

AT1G63070 N/A N/A N/A N/A

AT1G63080 N/A N/A N/A N/A

AT1G63130 N/A N/A N/A N/A

AT1G63150 N/A N/A N/A 0.40

AT1G63360 N/A N/A N/A N/A

AT1G63400 1.24 N/A N/A N/A

AT1G63630 1.24 N/A N/A N/A

AT1G64100 N/A N/A N/A N/A

AT1G64580 N/A N/A N/A N/A

AT1G66370 N/A N/A N/A N/A

AT1G66690 N/A N/A N/A N/A

Continued 69 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT1G66700 N/A N/A N/A N/A miRNA targets AT1G66720 N/A N/A N/A N/A

AT1G67450 N/A N/A N/A N/A

AT1G69170 0.73 N/A N/A N/A

AT1G69770 N/A N/A N/A N/A

AT1G72830 -0.57 N/A N/A N/A

AT1G76810 N/A N/A N/A N/A

AT1G77405 N/A N/A N/A N/A

AT1G77850 N/A N/A N/A N/A

AT2G04920 N/A N/A N/A N/A

AT2G17830 N/A N/A N/A N/A

AT2G18780 N/A N/A N/A 1.10

AT2G22740 N/A N/A N/A 0.80

AT2G22840 0.95 N/A N/A N/A

AT2G25980 -1.93 N/A N/A -1.30

AT2G26950 N/A N/A N/A N/A

AT2G27520 N/A N/A N/A N/A

AT2G28190 N/A N/A N/A N/A

AT2G28350 N/A N/A N/A -1.10

AT2G28550 -0.74 N/A N/A -4.30

AT2G29130 -4.38 N/A N/A -0.60

AT2G30210 -2.36 N/A N/A -3.40

AT2G31070 N/A 0.80 N/A N/A

AT2G32460 N/A N/A N/A N/A

AT2G33810 -2.58 N/A N/A N/A

AT2G34710 N/A 0.80 0.90 0.90

Continued 70 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT2G35160 N/A N/A N/A 0.70 miRNA targets AT2G36400 N/A N/A N/A N/A

AT2G38080 -1.44 N/A N/A N/A

AT2G39250 N/A N/A N/A N/A

AT2G41720 N/A N/A N/A 2.30

AT2G42200 N/A N/A N/A N/A

AT2G45160 N/A N/A N/A N/A

AT2G47460 N/A N/A N/A -3.80

AT3G03580 N/A N/A N/A N/A

AT3G05690 N/A N/A N/A N/A

AT3G08500 N/A N/A N/A N/A

AT3G09220 -4.02 -1.38 -1.04 -1.10

AT3G11440 N/A N/A N/A N/A

AT3G13820 N/A N/A N/A N/A

AT3G13830 N/A N/A N/A N/A

AT3G15030 N/A N/A N/A N/A

AT3G15170 N/A N/A N/A N/A

AT3G15270 N/A N/A N/A N/A

AT3G15640 N/A N/A N/A 1.00

AT3G16710 N/A N/A N/A N/A

AT3G16880 N/A N/A N/A N/A

AT3G17280 N/A N/A N/A N/A

AT3G17490 N/A N/A N/A N/A

AT3G19880 N/A N/A N/A N/A

AT3G19890 N/A N/A N/A N/A

AT3G20710 N/A N/A N/A N/A

Continued 71 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT3G20910 N/A N/A N/A N/A miRNA targets AT3G21170 N/A N/A N/A N/A

AT3G22350 N/A N/A N/A N/A

AT3G22700 N/A N/A N/A N/A

AT3G22710 N/A N/A N/A N/A

AT3G22890 N/A N/A -0.64 N/A

AT3G23690 N/A N/A N/A 1.10

AT3G26810 N/A N/A N/A -1.50

AT3G44860 -5.71 N/A N/A N/A

AT3G45090 0.68 N/A N/A N/A

AT3G49510 N/A N/A N/A N/A

AT3G49520 N/A N/A N/A N/A

AT3G54990 N/A N/A N/A -1.50

AT3G57230 N/A N/A N/A N/A

AT3G57920 N/A N/A N/A N/A

AT3G60630 N/A N/A N/A N/A

AT3G62980 N/A N/A N/A -0.70

AT4G00150 N/A N/A N/A N/A

AT4G03190 N/A 1.78 1.97 N/A

AT4G08990 N/A N/A N/A N/A

AT4G10780 N/A N/A N/A N/A

AT4G14680 N/A N/A N/A 3.40

AT4G18390 N/A N/A N/A N/A

AT4G19440 1.75 N/A N/A N/A

AT4G24150 -2.37 N/A N/A N/A

AT4G26800 N/A N/A N/A N/A

Continued 72 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT4G26930 N/A -0.77 -0.74 N/A miRNA targets AT4G27190 N/A N/A N/A N/A

AT4G30080 N/A N/A N/A N/A

AT4G32880 1.06 N/A N/A N/A

AT4G33290 -3.63 N/A N/A N/A

AT4G36920 0.63 N/A N/A -2.50

AT4G37740 N/A N/A N/A N/A

AT5G05400 N/A N/A N/A -2.50

AT5G06100 N/A N/A N/A N/A

AT5G06510 N/A N/A N/A N/A

AT5G07680 1.09 -0.98 N/A -5.70

AT5G10180 N/A 0.91 0.85 2.30

AT5G12840 1.56 N/A N/A N/A

AT5G16640 N/A N/A N/A N/A

AT5G23480 1.54 N/A N/A N/A

AT5G28520 N/A -0.73 -1.20 N/A

AT5G36190 N/A N/A N/A N/A

AT5G37020 N/A N/A N/A N/A

AT5G38550 N/A N/A N/A N/A

AT5G39610 N/A -1.42 -1.30 -4.00

AT5G41170 0.84 N/A N/A N/A

AT5G41610 2.27 N/A N/A 2.30

AT5G43270 N/A N/A N/A N/A

AT5G43730 N/A N/A N/A -2.10

AT5G43740 N/A N/A N/A N/A

AT5G43780 -1.75 -1.40 -1.30 -4.50

Continued 73 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT5G46680 N/A N/A N/A N/A miRNA targets AT5G47260 N/A N/A N/A N/A

AT5G49870 N/A N/A N/A -0.60

AT5G50570 N/A N/A N/A N/A

AT5G50670 N/A N/A N/A N/A

AT5G51270 N/A N/A N/A N/A

AT5G53660 N/A N/A N/A N/A

AT5G53890 N/A N/A N/A N/A

AT5G53950 N/A N/A N/A N/A

AT5G55020 N/A N/A N/A N/A

AT5G57250 N/A N/A N/A N/A

AT5G60020 N/A N/A N/A -3.00

AT5G60120 0.76 N/A N/A -2.60

AT5G60690 N/A N/A N/A N/A

AT5G60760 1.14 N/A N/A -3.00

AT5G61430 -0.17 N/A N/A -1.20

AT5G62310 N/A N/A N/A N/A

AT5G65560 N/A N/A N/A N/A

AT5G67180 N/A N/A N/A 0.80

AT1G03560 N/A N/A N/A N/A ta-siRNA targets AT1G12620 N/A N/A N/A N/A

AT1G12700 N/A N/A N/A N/A

AT1G12775 N/A N/A N/A 0.80

AT1G31840 N/A N/A N/A N/A

AT1G51670 N/A N/A N/A N/A

AT1G54260 2.61 N/A N/A N/A

Continued 74 Table 2.1 continued

RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT1G56650 N/A N/A N/A N/A ta-siRNA targets AT1G62590 1.74 N/A N/A N/A

AT1G62680 N/A N/A N/A N/A

AT1G62720 1.70 N/A N/A N/A

AT1G62860 3.04 N/A N/A N/A

AT1G62930 1.07 N/A N/A N/A

AT1G63070 0.40 N/A N/A N/A

AT1G63080 N/A N/A N/A N/A

AT1G63150 N/A N/A N/A 0.40

AT1G63400 N/A N/A N/A N/A

AT1G63630 N/A N/A N/A N/A

AT1G64100 3.36 N/A N/A N/A

AT1G64310 0.57 N/A N/A N/A

AT1G64580 N/A N/A N/A N/A

AT1G66370 N/A N/A N/A N/A

AT1G66390 N/A N/A N/A N/A

AT1G74900 N/A N/A N/A N/A

AT2G17140 N/A N/A N/A N/A

AT2G33860 2.78 N/A N/A 1.30

AT4G21170 0.95 N/A N/A N/A

AT4G26800 1.99

AT4G28010 1.34 N/A N/A N/A

AT4G29770 N/A N/A N/A 3.50

AT5G16640 1.32 N/A N/A N/A

AT5G18040 1.49 N/A N/A 2.30

AT5G41170 N/A N/A N/A N/A

Continued 75 Table 2.1 continued RNAi 21dpi Syncytium Gene ID 14dpi gall 21dpi gall category giant cell 5 & 15dpi

AT5G60450 0.31 N/A N/A 0.80 ta-siRNA targets AT5G62000 0.91 N/A N/A -0.80

Table 2.1. Disruption of RNAi-related gene expression observed during nematode

infection. Log2 fold changes in feeding site or infected treatment versus non-feeding site or uninfected control are presented with a false discovery rate of < 5% for giant cell (Morse et al., 2010) and syncytium studies (Szakasits et al., 2009) and P < 0.05 after Bonferroni correction for the gall study (Jammes et al., 2005). N/A are not differentially expressed at the significance cutoff, and black cells are not represented on the CATMA chip. Among significantly differentially expressed genes, the number of upregulated RNAi machinery genes deviate significantly from an expected ratio of 1:1 in laser- capture microdissected giant cells (Chi-square test, p < 0.0005), galls at both 14 and 21 days post infection in comparison to uninfected tissue (Chi-square test, p < 0.1), and in microaspirated syncytia (Chi-square test, p < 0.001). Among significantly differentially expressed miRNA-targeted genes in giant cells, the majority are upregulated (Chi-square test, p < 0.1). A disproportionate number of ta-siRNA-targeted genes are also upregulated in both giant cell and syncytium studies (Chi-square test, p < 0.05).

76

CHAPTER 3: THE MATURE FEMALE ROOT-KNOT NEMATODE TRANSCRIPTOME

3.1. ABSTRACT

The southern root-knot nematode (RKN, Meloidogyne incognita Chitwood) is the plant-parasitic nematode most economically damaging to agricultural production globally. The strategies currently utilized to control RKN infestations include host resistance, which is not available in all crops, nematicides that are increasingly coming under tighter restrictions for use, and biological controls, which are still lagging in providing consistent nematode control. Proteinaceous toxins disrupting nematode digestion and small RNAs targeting nematode transcripts for nematode uptake have also met with varying success in laboratory and field studies. While transcriptional and proteomic approaches have predominantly focused on early stages of infection, our study aims to provide more insight into strategies employed by adult female RKN. Through an RNA-Seq analysis of total RNA isolated from individually harvested female RKN, we have generated a whole transcriptome and putative secretome for this important life stage. For a subset of genes, expression was verified with RT-PCR in eggs and infective juveniles in addition to females. Transcriptomic analysis revealed gene expression patterns expected of this life stage, including the production of digestive enzymes and eggs. Interesting trends in expression potentially relevant to the parasitic interaction were observed among cell wall modifying proteins, proteases, as well as previously characterized effectors. Results have identified new targets for functional analysis to determine their role in this parasitic interaction.

77 3.2. INTRODUCTION

Root-knot nematodes (RKN, Meloidogyne spp.) are the most economically damaging plant-parasitic nematode, costing billions of dollars in damage to agricultural crops around the world every year (Nicol et al., 2011). RKN are known to parasitize over 2,000 different plant species (Sasser, 1980), encompassing virtually every agricultural crop. A major species, which accounts for the most losses worldwide, is the southern root-knot nematode, Meloidogyne incognita. Sustaining their ability to cause widespread damage is the fact that they are notoriously difficult to control. Many chemical nematicides, while effective, are costly and face increasing restrictions due to their human and environmental health hazards. Research into the formulation and use of biological controls is increasing but this method still faces major obstacles with achieving both consistency and efficacy in the field. When available, host resistance is undoubtedly the favored method of control. Resistance conferred by the Mi gene is present in almost all commercially available tomato varieties (Milligan et al., 1998; Roberts and Thomason, 1986). Although very effective in many cases, resistance-breaking strains continue to develop (Jacquet et al., 2005; Williamson, 1998). For many crops, effective resistance to RKN is unavailable. Genetic engineering is offering new approaches; for example, expression of small RNA molecules that target and silence nematode transcripts (Rosso et al., 2009), or expression of proteinaceous toxins (Urwin et al., 1997a), but success has yet to be realized in the field. The shortcomings of these control strategies warrant further investigation into this parasitic interaction in order to define new targets, deliver toxins more directly, or develop entirely new strategies. There are several aspects of the obligately biotrophic M. incognita lifecycle that make it a particularly difficult pathogen to control. The nematode eggshell is capable of preventing the entry of many small molecules, thus protecting the developing juvenile from surrounding microbiota (Edgar et al., 1994). Once developed, the infective second- stage juvenile (J2) hatches and must locate a host root. When a root is located, the J2 secretes an assortment of enzymes and uses its stylet (a spear-like mouthpart) to penetrate and migrate intercellularly within the root. After initiating a feeding site in the vascular 78 cylinder, the J2 undergoes three additional molts. Four to seven cells are selectively probed by the nematode stylet through which several more molecules, including a diverse array of proteins, some mimicking plant peptides and hormones, are secreted (Mitchum et al., 2013). These secreted products are presumed to be largely responsible for the drastic reprogramming of those root cells into the feeding site composed of “giant cells.” The selected cells undergo multiple rounds of endoreduplication, resulting in multinucleated cells roughly 100 times their original size (de Almeida Engler et al., 1999). Giant cell characteristics include a proliferation of organelles, thickened and highly invaginated cell walls, and high metabolic activity, essentially generating a nutrient sink for the female nematode (Caillaud et al., 2008). The successful initiation of a feeding site by a J2 is followed by loss of most of its somatic musculature, rendering the nematode immobile for the remainder of its life. The adult female M. incognita relies upon successful maintenance of the giant cells in order to acquire the energy and nutrition necessary for egg production. This total reliance on a handful of cells makes it a stage of interest for development of new strategies for control. Although a great deal has been learned from research focused on this parasitic interaction, including identification and characterization of many of the secretions M. incognita utilize to successfully parasitize their host, the strategies used by the adult females to maintain the relationship throughout egg production remain underexplored. Genome sequencing has enabled enormous advances in our understanding of how plant parasitism in the phylum Nematoda has evolved (Abad et al., 2008). Adaptation to plant parasitism has involved the incorporation of bacterial genes via horizontal gene transfer, including a suite of cell wall-degrading enzymes, other enzymes thought to aide in the evasion of host defenses such as chorismate mutases and a cyanate lyase, and an invertase likely assisting in nutrient acquisition (Danchin et al., 2010; Haegeman et al., 2011; Whiteman and Gloss, 2010). Some gene sets appear to have expanded, based on the presence of InterPro domains, such as C48 SUMO (small ubiquitin-like modifier) family cysteine proteases, whereas others appear to have contracted, such as those involved in chemosensory activity. Some of the changes in gene abundances could be attributed to the sedentary endoparasitic life history of RKN. For the 79 majority of its lifecycle, the nematode is surrounded by host tissue, thus increasing its need to evade host defense responses, but decreasing its need for chemoreceptors to sense its environment. Genomics has supplied us with information regarding the capacity of M. incognita to parasitize plants; while proteomic and transcriptomic studies have also allowed researchers to identify several genes involved in parasitism, some of which have been characterized for their role in this parasitic interaction (Bellafiore et al., 2008; Mitchum et al., 2013). The obligate nature of M. incognita parasitism makes it particularly tricky to study, especially during later stages of infection. The nematodes’ esophageal glands are the primary source of parasitism proteins, referred to as effectors (Mitchum et al., 2013). Due to this, researchers have focused on stylet secretions, either chemically induced (Hussey et al., 1994) or directly microaspirated contents of the dorsal and subventral esophageal glands (Huang et al., 2003). Several putative effector genes have been identified with transcriptomic studies (Davis et al., 2000; Davis et al., 2008; Haegeman et al., 2011; Hewezi and Baum, 2013b). A diverse array of effectors, from cell wall modifiers to plant signal peptide mimics have been identified through sequence analyses and, in some cases, confirmed via functional analyses (Mitchum et al., 2013; Smant et al., 1998; Wang et al., 2005). The majority of putative effectors remain both uncharacterized and lack any homology to known proteins; these novel proteins are referred to in the literature as “pioneers.” Isolating gland cell-specific RNA is somewhat problematic in the adult female, and isolating stylet secretions impossible, because the body is delicately buried in root tissue. These techniques also are unable to detect other important secretions not originating from the esophageal glands, such as those originating from the hypodermis and amphids. We used a whole transcriptome analysis of isolated adult female M. incognita to address the obstacles and limitations of these other approaches. Further analysis of the putatively secreted set of proteins is still warranted, as many in this category are widely recognized for their roles in pathogenesis (Vanholme et al., 2004). Other studies on M. incognita have focused on early life stages and on gland cell-specific expression or secretion. In order to better understand the adult female life stage, we provide a 80 comprehensive investigation of the adult female transcriptome and putative secretome as it relates to parasite biology and potential giant cell maintenance strategies.

3.3. MATERIALS AND METHODS

3.3.A. Nematode colony maintenance and extraction The RKN (M. incognita) population was maintained in dixenic culture on Arabidopsis (ecotype Columbia-0) as previously described (Hammes et al., 2005). Six weeks after infestation with 1,000 RKN eggs, adult females were excised from root tissues with dental picks and placed in TBS buffer (TRIS-buffered saline, pH 8) on ice. Females were rinsed with TBS buffer over a sieve, placed in a new Eppendorf tube and treated with RNase A (Promega, Madison, WI) according to the manufacturer’s protocol. After 15 minutes of incubation at 37°C with RNase, excess buffer was removed and the sample was processed for RNA isolation. Eggs were also harvested from dixenic Arabidopsis culture plates. Infested roots from plates, six to seven weeks after infestation with 1,000 RKN eggs, were removed from the agar medium and eggs were isolated from the gelatinous matrices with sodium hypochlorite (Hussey and Barker, 1973). Eggs were centrifuged and concentrated into approximately 50 μl pellets. For pre-parasitic second stage juvenile (J2) samples, additional eggs were isolated as described, placed on a 25 μm nylon mesh, saturated with water, and allowed to hatch. After three days J2 were similarly concentrated into approximately 50 μl pellets. Egg and J2 samples were then subject to RNA isolation consistent with the method used for adult females.

3.3.B. RNA isolation and Illumina library synthesis Total RNA was extracted from three independent pools of 150 adult female RKN from independent batches of Arabidopsis culture plates with the RNeasy Plant Mini Kit protocol (Qiagen, Valencia, CA). Homogenization of the tissue was performed with an eppendorf pestle in RTL buffer (10% β-mercapto-ethanol). RNA quantity and quality were assessed with the Qubit® 2.0 Fluorometer (Life Technologies, Carlsbad, CA) and a

81 Bioanalyzer 2100 (Agilent Technologies, Palo Alto, CA) respectively. All samples selected for cDNA synthesis had RNA integrity numbers (RIN) ≥ 7. RNA from each pool (1.5 μg) was used to generate an adaptor-tagged double- stranded DNA library for RNA-Seq with the TruSeq Stranded mRNA Sample Prep LS protocol (Illumina, San Diego, CA) following the manufacturer’s protocol. Quantification of DNA was done with the Qubit® 2.0 Fluorometer (Life Technologies, Carlsbad, CA) and quality was assessed with the Bioanalyzer (Agilent Technologies, Palo Alto, CA). The pooled samples were diluted to 4 nM and combined to generate one multiplexed DNA library for each replicate.

3.3.C. Illumina sequencing and transcriptome assembly The cDNA library (10 pmoles) was sequenced on one flow cell lane with the Illumina HiSeqTM2000 platform at the Ohio State University Comprehensive Cancer Center (OSUCCC). Both ends of the library were sequenced to generate 100 nt raw paired-end reads. The Illumina Analysis Package CASAVA 1.8.2 was used to perform bcl conversion and demultiplexing. Image deconvolution and quality value calculations were carried out with the Illumina GA pipeline v1.8. The RKN raw reads were imported into CLC Genomics Workbench (v6.5.1, CLC Bio) and trimmed for quality, adapter indexes and poly(A) tails with the default settings (Ambiguous limit = 2, quality limit = 0.05). Processed reads were assembled de novo by means of two independent approaches. First, with the CLC Bio algorithm based on de Bruijn graphs and the optimized parameters [Word Size = 54, Bubble Size = 650, Length Fraction = 0.6, Similarity Fraction = 0.95] contigs of ≥250 nt were assembled; and second, with the Oases v0.2.08 (Schulz et al., 2012) with Kmer sizes of 53, 59, 65, 71, 77, 83, and 89. Transcripts with ≥90% sequence similarity were collapsed into clusters and the longest read retrieved using CD-HIT-EST (Li and Godzik, 2006). The assemblies were merged into a final assembly with Minimus2 (Sommer et al., 2007). The raw sequence reads were submitted to the NCBI short sequence read archive under the accession number SRXXXXXXXX.

82 De novo assembled transcripts were assigned hierarchical gene ontologies (GO terms) on the basis of biological processes, molecular functions, and cellular components with the platform-independent Java™ 6 implementation of the BLAST2GO software (Gotz et al., 2008). The top five tBLASTx hits to the nr database with a cut-off E-value of 10-3 were considered for GO annotation.

3.3.D. RKN secretome analyses The six open reading frame (ORF) amino acid sequences were predicted from the transcript sequences of RKN with ORF-Predictor (Min et al., 2005). Only the subset of predicted sequences ≥50 amino acids was used in subsequent analyses. The SignalP 4.1 neural networks algorithm (Petersen et al., 2011) was implemented to detect putative transmembrane proteins with signal peptide secretion and cleavage site signatures in their amino acid sequences with the default settings for D-score. For the subset of predicted secreted proteins, gene ontology was performed with BLAST2GO software (Gotz et al., 2008) for further functional annotation based on the protein sequences. The top five BLASTp matches to the nr database with a cut-off E- value of 10-3 were considered for hierarchical GO characterization. In addition, desktop downloaded tBLASTx software (E-value <10-10) was used to perform pair-wise comparisons between the RKN sequences and four other nematode databases: the EST database of Meloidogyne hapla (northern root-knot nematode, order Tylenchida, 24,453 sequences, accessed from NCBI Genomes FTP); the cDNA database of Caenorhabditis elegans (soil nematode, order , 32,201 sequences accessed from NCBI Genomes FTP); the cDNA database of Teladorsagia circumcincta (order Strongylida, 27,593 sequences accessed from Nematode.net); and Dictyocaulus viviparus (bovine lungworm, order Strongylida, 36,626 sequences accessed from Nematode.net). For the subset of predicted secreted proteins of each nematode, BLAST2GO was used to identify proteins that had a significant BLASTp hit to the nr database (E-value <10-10). In addition, desktop downloaded tBLASTx software (E-value <10-10) was used to carry out pair-wise comparisons to the NCBI EST database of the northern root-knot nematode, M. hapla (24,453 sequences, accessed June 15, 2014).

83

3.3.E. Identification of plant material in RKN In order to query our RKN transcriptome for transcript sequences of plant origin, preprocessed reads were mapped to the Arabidopsis thaliana cDNA database (33,602 sequences, accessed from TAIR) with the CLC Bio RNA-Seq analysis function and stringent settings of Length Fraction = 0.95 and Similarity Fraction = 0.99 (default settings herein). A. thaliana sequences containing a minimum of 5 mapped unique reads in all three samples were putatively determined to be present in RKN. As these transcripts were presumably ingested, primers were designed to amplify multiple lengths of identified transcripts to verify their presence with reverse transcriptase PCR (RT-PCR) (PCR conditions described in section 3.3.F).

3.3.F. Reverse transcriptase PCR for expression validation. RNA isolation for RT-PCR experiments was done in the same manner as described above, except only 50 females were isolated per sample. RNA template (250ng) was treated with RQ1 DNase (Promega, Madison, WI) and used for cDNA synthesis with the GoScriptTM Reverse Transcription System (Promega, Madison, WI) following the manufacturer’s protocol and primed with oligo(dT)15. During cDNA synthesis a no-RT reaction was included for each RNA sample for detection of contaminating genomic DNA. cDNA template was diluted 2:1 and amplified with Phusion® High Fidelity DNA Polymerase (New England Biolabs, Ipswitch, MA). Oligonucleotides were synthesized by Integrated DNA Technologies (Coralville, IA, U.S.A.); primer sequences are listed in tables 3.1 and 3.2. The PCR cycle conditions used were: initial denaturation at 98°C for 30 seconds; 28 or 35 cycles of 98°C for 30 seconds, 57°C for 15 seconds, and 72°C for 15 seconds; and a final extension at 72°C for 10 minutes.

84

3.4. RESULTS

3.4.A. Sequence assembly In these experiments, we performed RNA-Seq analysis on three independent replicates of 150 adult females of the southern root-knot nematode (RKN; Meloidogyne incognita) isolated six weeks after infesting Arabidopsis seedlings with eggs. Adult females with no or few eggs were chosen to minimize the abundance of transcripts related to embryogenesis and thus maximize the relative abundance of those transcripts pertinent to the interaction. Following total RNA extraction and library synthesis, the pooled cDNA libraries were sequenced, which generated 136,183,194 paired-end, 100 nt reads. After trimming (quality, adapters, poly(A) sequences), ~114.7 million reads were obtained (85.4 M reads in pairs). The de novo assembled transcriptome consisted of 28,106 non-redundant M. incognita transcripts (≥226 nt; x̅ = 762 nt). Functional characterization of the contigs with tBLASTx indicated 10,200 (36%) had a significant match to the non-redundant (nr) database (Fig. 3.1.). The most common ortholog matches were to Caenorhabditis spp. (14%), Loa loa (13%), Brugia malayi (11%), and Strongyloides papillosus (11%). Plant-parasitic nematodes represented 4.8% of transcripts with a significant BLASTx hit (E value < 10-3) to the nr database, among which Meloidogyne spp. made up the majority of top hits.

3.4.B. The M. incognita secretome Where possible, transcript sequences were translated into their putative amino acid sequences. A total of 24,694 M. incognita sequences of ≥50 amino acids were identified. In silico analysis of transcripts coding for secreted proteins with SignalP (Nordahl et al., 2011) revealed 1,696 M. incognita sequences (6.9%) had signatures for signal peptide secretion and cleavage sites. Furthermore, 8.6% of the putative secreted proteins had a transmembrane domain signature.

85 For the subset of putatively secreted proteins, 528 (31%) and 782 (46%) M. incognita had significant tBLASTx and BLASTp matches to the nr database, respectively. BLAST results were compiled, resulting in 833 (49%) putatively secreted proteins with annotation data. Gene ontology (GO) annotation was done to categorize transcripts into their putative biological process, cellular compartment, and molecular function. Functional categorization of this set of genes resulted in 192 GO terms describing 329 contigs in the sub-ontology “biological process.” The top twenty common GO terms represent approximately 40% of genes encoding putatively secreted proteins with associated GO terms (Fig. 3.2). The top twenty terms for the sub-ontology “cellular compartment” represent almost 90% of this set of genes, with the most abundant terms predictably being associated with membrane systems and extracellular space (Fig. 3.3). Just over 50% of genes with associated GO terms were represented by the top twenty terms in the sub-ontology “molecular function,” which includes functions indicative of the parasitic interaction and digestion (Fig. 3.4). InterPro software (Mitchell et al., 2015) was used to determine the presence of functional domains within our in silico translated transcriptome. Many of the most abundant transcripts, based on average coverage and paired-end reads mapped, have domains expected for this life stage, including lipid transport and vitellinogen (an egg yolk precursor) (Table 3.3.). The most abundant domains in the subset of highly expressed putatively secreted proteins include various protease activities known to be involved in digestion and the parasitic interaction (Table 3.4.).

3.4.C. Comparison of nematode secretomes Pairwise sequence comparisons were carried out between the 1,696 M. incognita putatively secreted proteins and four other nematode species with available EST or cDNA databases: Meloidogyne hapla; the free-living Caenorhabditis elegans; and the animal parasitic nematodes Teladorsagia circumcincta and Dictyocaulus viviparous. Nearly half (819) of the M. incognita sequences had a significant match to at least one of the nematode databases and 7.7% (130) matched all four species (E value <10-10) (Fig. 3.5). As expected, M. incognita had the largest number of total matches (568) and unique

86 matches (284) to the most closely related species, M. hapla. M. incognita had the second most matches to C. elegans (507) and the least number to T. circumcincta (224), which likely partially reflects the quality and completeness of those databases in addition to their specialized parasitic niches.

3.4.D. Analysis of specific gene groups Expression of genes of interest was verified with RT-PCR, including additional life stages for relative expression comparisons with eggs and pre-parasitic J2 (Table 3.5.). The expression of select cell wall modifiers, proteases, known virulence factors, and gland-specific transcripts were analyzed across these growth stages. M. incognita secrete a battery of cell wall modifying enzymes during early stages of infection, allowing them to penetrate their host root and migrate intercellularly before initiating a feeding site. Expression of similar cell wall modifying enzymes is also present in our adult female dataset, which includes forty contigs with a cell wall-modifier top BLASTx hit. The expression of a β-1, 4-endoglucanase was verified with RT-PCR and was also observed in eggs and in J2 (Fig. 3.6.H). Interestingly, when comparing relative expression between life stages, a pectate lyase, which often is associated with penetration and intercellular migration, appears to be upregulated in the adult female (Fig. 3.6.B) as well. An expansin in our dataset also appears to be upregulated at this later phase, but with no apparent expression in eggs or J2 (Fig. 3.7.C). Forty-one transcripts with a cell wall-degrading or modifying top BLASTx hit are present in our female M. incognita transcriptome (Table 3.6). The expression of other proteases was also analyzed. Cysteine proteases were represented in ten top BLASTx hits and among the most abundantly expressed putatively secreted set. RT-PCR validation of a chymotrypsin gene expression (AY714229) was consistent with that previously reported (de Souza Júnior et al., 2013), with moderate expression in both eggs and females and none detected in pre-parasitic J2 (Fig. 3.7.B). Also in the family of serine proteases, three transcripts identified in the S16 Lon protease subfamily (IPR008269), which represents one of the subfamilies that has expanded in the M. incognita genome versus the free-living C. elegans. Our analysis of a reported

87 cathepsin-D aspartic protease (Mi-asp1; DQ360827) was also consistent with a previous study (da Rocha Fragoso et al., 2009), present in all life stages and elevated in the adult female (Fig. 3.7.E). A transcript with homology to cathepsin L (Mi-cpl-1) was also present in our female dataset (Neveu et al., 2003). Other genes of note were six metalloproteases, which are not commonly found in plant-parasitic nematodes, but have been discovered in the potato cyst nematode, Globodera rostochiensis, and RKN secretions (Bellafiore et al., 2008). Other transcripts of interest were originally isolated from RKN esophageal glands, characterized, and found to play an important role in the success of the parasitic interaction. Chorismate mutase (CM), first identified in the subventral glands of M. javanica (Lambert et al., 1999), is a key enzyme in the biosynthesis of aromatic amino acids, and therefore may help subvert host tyrosine-dependent lignification or other plant defense responses. In addition to the CM that has been characterized (AY422834; M. incognita) we also observed and confirmed the expression of another CM (DQ222223; M. arenaria), which appears to increase in expression at the J2 and adult female stage (Fig. 3.6.E). Calreticulin is another transcript of interest as it has been found to play a role in animal parasitism (Nakhasi et al., 1998) as well as suppress basal plant defenses (Jaouannet et al., 2013). Expression in M. incognita was observed in the subventral glands of parasitic juveniles and the dorsal gland of adult females (Jaouannet et al., 2013; Jaubert et al., 2002). We observed calreticulin expression in eggs and females, but to a lesser extent in pre-parasitic J2 (Fig. 3.8.D). The importance of calreticulin to the parasitic interaction has also been verified. When its expression was knocked-down in J2 with dsRNA, virulence was reduced (Dubreuil et al., 2009), and knockdowns on transgenic Arabidopsis also resulted in a reduction in virulence following root penetration (Jaubert et al., 2005). The majority of M. incognita secreted proteins have unknown function and even lack homology to characterized domains; these are often referred to as “pioneer” genes. Of twenty-seven previously reported pioneer genes expressed in the esophageal glands of life stages from J2 to J4-adult (Huang et al., 2003), nineteen were found in our adult females (Table 3.7.). One such pioneer, gland protein 23 (M. incognita; Minc18502) 88 (AY134442), is expressed in the dorsal gland of parasitic juveniles and adults but was not detected in pre-parasitic juveniles (Huang et al., 2003). The gland protein 23-like transcript in our dataset was present in eggs, females, and pre-parasitic juveniles, albeit to a lesser extent (Fig. 3.8.E). In addition to awaiting functional characterization, some proteins that play a role in this parasitic interaction are expressed in tissues other than the esophageal glands. For example, the amphids were found to express factors important to the interaction (Semblat et al., 2001). A transcript in our adult females shared homology to an amphid-secreted protein in the cyst nematode, Globodera rostochiensis (Jones et al., 2000). Consistent with findings in the cyst system, we found this transcript to be expressed in all life stages tested (Fig. 3.6.G). Other genes of note, and among the most abundant transcripts, were a vitellogenin and a mucin (Table 3.3). Vitellogenin-6 is secreted from the intestine during egg laying (Nakamura et al., 1999); it has metal-binding capacity and is thus hypothesized to protect cells from oxidation (Ishii et al., 2002). As expected, analysis of vitellogenin expression was absent in egg and J2 samples and abundant in adult females (Fig. 3.6.C). Any role of mucin in the parasitic interaction is not known in plant-parasitic nematodes. Homologues in the animal parasite, T. canis, provide variability to its surface coat thus contributes to evading host immunity (Tetteh et al., 1999). Mucin proteins may play a role in host specificity of P. penetrans endospore attachment to the root-knot nematode cuticle (Davies, 2009). The expression of a putatively secreted patatin was also analyzed and found to be present in all stages tested (Fig. 3.6.D). Patatin is a non-specific lipid acyl hydrolase, which has displayed nematistactic activity when expressed in plants (Burrows and De Waele, 1997). Endogenous expression in plants has been found to be responsive to stress; interestingly, the Arabidopsis patatin (At2g26560) is significantly downregulated during RKN-induced gall formation (Jammes et al., 2005).

3.4.E. Plant transcript discovery and analysis In addition to M. incognita transcripts, this dataset also indicated the presence of transcripts of host plant origin within the whole-nematode RNA samples. Those transcripts that were consistently detected across three independent biological replicates 89 of RNA-Seq are summarized in Table 3.8. As an independent method of detection, additional M. incognita female samples were isolated and Arabidopsis transcripts were amplified via RT-PCR. Two portions of the Catalase 3 (AT1G20620; 219bp and 476bp) and the Translationally Controlled Tumor Protein (TCTP) transcripts (AT3G16640; 249bp and 640bp) were amplified in two independent samples (Fig. 3.9).

3.5. DISCUSSION

A great deal has been learned from the genome sequence of M. incognita with regards to the M. incognita genetic adaptation for parasitism (Abad et al., 2008). The general aim of this study was to examine the transcriptional profile of female RKN, which represents a phase of the parasitic interaction that is less understood. The transcriptional as well as putative secretion profile at this time point gives us some insight into the differential needs of female M. incognita compared with earlier stages, which must maintain their feeding sites but do not have to meet the metabolic needs of reproduction. We examined approximately 24,694 M. incognita expressed sequences, some of which may reflect splice variants from identical genes or PCR artifacts, as the number exceeds the estimated 19,212 protein-coding genes in the genome (Abad et al., 2008). Of the expressed sequences in my study, there were 1,696 determined to be putatively secreted, roughly 40% in comparison to the 4,250 predicted in the genome, which may reflect different cutoff settings (measured by Mathews Correlation Coefficient); thus, sensitivity for signal peptide calling. It is also likely that some transcripts may be absent from our secreted set due to fragmentation or cleavage of the signal peptide prior to sequencing or during assembly. Most M. incognita genes involved in parasitism are thought to reside in this secreted set, as they have the potential to interact at the interface of the interaction as well as be secreted directly into the host cells that are serving as the nematode feeding site. The M. incognita genome encodes 339 proteases, 92 of which are putatively secreted (Abad et al., 2008). Parasitic nematode proteases, plant and animal-parasitic

90 alike, are known to perform a variety of functions that contribute to virulence. Some proteases degrade host proteins such as components of the plant cell wall or cellular components prior to uptake and digestion, and some inhibit defense pathways (Tort et al., 1999). Proteases are also secreted in the nematode intestine for digestion and are a popular target for antihelminthics (Urwin et al., 1997b). Cysteine proteases in M. incognita are known to be important in digestion (Koritsas and Atkinson, 1994) and were among the most abundantly represented protease type in our female dataset. While it is likely many of these cysteine proteases are secreted into the intestine for digestive purposes, a potential alternative role in parasitism would need to be experimentally determined. Are any of the proteases expressed during this stage localized to the esophageal glands? An aspartyl protease-like protein (Mi-ASP2), for instance, is secreted into the plant apoplasm from the nematode’s subventral glands (Vieira et al., 2011). Other proteases expressed during the sedentary phase may be involved in pre-digestion of nutrients prior to uptake, or may have a role in inhibiting defense responses as was demonstrated for cysteine proteases secreted by the bacterium, Pseudomonas syringae (Dowen et al., 2009). Infective juveniles secrete an assortment of cell wall-degrading enzymes to penetrate their host root and migrate between cells during the initial stage of infection (Hewezi and Baum, 2013a; Wieczorek et al., 2014). β-1, 4-endoglucanase was the first cell wall-degrading enzyme discovered in cyst nematode secretions (Smant et al., 1998), and it has since been found in several other plant-parasitic nematode secretions (Haegeman et al., 2012). For example, Mi-eng-2 was characterized for its role in cell wall degradation and expression was confirmed in eggs, J2, and adult females (Ledger et al., 2006); these expression results were confirmed in our study. Cell wall-degrading, carbohydrate-active enzymes (CWDE) represent a remarkable nematode adaptation to plant-parasitism, as they are unknown in other metazoans. Sixty-one plant CWDE are putatively encoded in the genome: 21 cellulases, six xylanases, two polygalacturonases, 30 pectate lyases, and two arabinases, as well as two invertases and 20 candidate expansins involved in cell wall modifying processes (Abad et al., 2008). The 40 CDWE- like or cell wall-modifying top BLASTx hits in our dataset included: one cellulase, seven 91 xylanases, two polygalacturonases, five pectate lyases, eight β-1, 4-endoglucanases, as well as 15 expansins and one cellulose binding protein. In addition to cell wall-degrading enzymes, M. incognita secrete other factors that interact with the host cell walls but have no hydrolytic activity. Expansins, for example, disrupt bonds between cell wall components, making them more accessible to hydrolytic enzymes and thus play an important role in cell expansion (McQueen-Mason and Cosgrove, 1995). The first nematode-encoded (indeed, first animal-encoded) expansin was discovered in Globodera rostochiensis (Qin et al., 2004a). Altered expression of host expansins have been observed in other interactions, such as H. schachtii in Arabidopsis (Wieczorek et al., 2006), and when the expression of expansins is knocked down in tomato, M. javanica reproduction and giant cell size are both reduced (Gal et al., 2006). Expansin-like proteins are presumably secreted into the plant apoplast during invasion to aid the nematode’s migration through root tissue (Qin et al., 2004b; Tomalova et al., 2012). The expression of an expansin examined in our study was apparent in adult females but absent in eggs and J2, indicating that this particular expansin may be involved in cell wall modification surrounding the feeding site, allowing for the rapid expansion of giant cells and/or the developing female, rather than the cell wall loosening that occurs during invasion. Another example is the cellulose binding protein, MI-CBP-1, which has been cloned and immunologically determined to be secreted from infective juveniles (Ding et al., 1998). Mi-cbp-1 also appears to be expressed in adult females in our dataset. The expression of some host pectate lyases also increases during maintenance and utilization of giant cells (Wieczorek et al., 2014), although mutant analyses showed that this increase in expression is more functionally relevant in the success of feeding sites of a different parasite, beet cyst nematode. It is possible that the RKN-expressed pectate lyase is somewhat redundant to this increase in host expression. It would be interesting to investigate whether these nematode pectate lyases that are expressed by the female nematode function to accommodate the expansion of both the giant cells and female body. Are any of these enzymes secreted from the rectal glands, indicating they may play a role in the emergence of the egg mass from the confines of the root? The expression of 92 several of these genes during the later stage of infection implies the continued importance of cell wall modifications during this stage of parasitism, potentially functioning in giant cell maintenance and expansion, and accommodating the expansion of the growing female and emergence of the developing egg mass from the root. In 2001, Semblat et al. reported on an M. incognita amphid-secreted avirulence factor (MAP-1). Amphids are chemosensory organs open to the environment located on lateral sides of the nematode’s head; amphids are known to secrete proteins (Perry, 1996). The amphid-secreted transcript present in our dataset (contig_13050) does not share significant sequence similarity at the amino acid level with MAP-1, but this finding highlights the possible importance of non-esophageal gland secretions during the parasitic interaction. Although esophageal glands have proven to be a treasure-trove of effectors, we would be remiss to disregard other secreted factors (e.g. amphidal, rectal, intestinal). For example, cathepsin L (Mi-cpl-1) is expressed in intestinal cells and thus likely involved in digestion (Neveu, 2003); it is still a worthy target, as RNAi knockdown reduced the number of developed females by approximately 60% (Shingles et al., 2007). Lastly, the possible presence of plant-derived RNA in adult females may provide some new insights into M. incognita feeding and giant cell anatomy. Previous reports have suggested that the feeding tube organelle located inside the giant cell in proximity to the stylet has a size exclusion limit of 40 kDa (Böckenhoff and Grundler, 1994). Transcripts of host origin of up to 872 nt were recovered from female M. incognita, which far exceeds the widely accepted exclusion limit at just under 300 kDa; however, the size of RNA molecules is greatly impacted by their secondary structure. Therefore, further studies to examine the parameters that affect passage through the nematode feeding tube will involve the expression of tagged proteins of known sizes and chemical properties. Further work will be needed to determine whether any of the specific host transcripts found in this study have a functional role in the interaction, or whether their abundance and/or subecullar localization in giant cells makes them likelier indirect targets for uptake.

93 Root-knot nematodes represent a serious threat to agricultural production around the globe. Effective management strategies are restricted due to this nematode’s vast host range and the limited availability of host resistance. New strategies involving the use of biological controls and genetic engineering show promise, but are not yet displaying consistent results in the field. Results from our study of the transcriptome of mature female M. incognita has pointed to new candidate genes involved in parasitism. Future experiments will be dsigned to investigate their potential role in the parasitic interaction by localizing their expression and determining the effects of the encoded proteins on host physiology and nematode susceptibility. Additionally, work pertaining to the parameters relevant to nematode ingestion may provide an opportunity to improve upon genetic engineering approaches by allowing researchers to increase the delivery of RNA molecules and proteinaceous toxins.

3.6. ACKNOWLEDGMENTS

We acknowledge Dr. Bryan Cassone for the bioinformatic analysis of RNA-Seq data, and Allison Grenell for female nematode isolation and plant transcript analyses. This work was supported in part by funding from The Ohio State University, Department of Plant Pathology and Center for Applied Plant Sciences, SoyRes Research Team.

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100 Table 3.1. Primer sequences used for detection of Arabidopsis transcripts in M. incognita cDNA libraries.

Amplicon Gene ID Forward primer (5’-3’) Reverse primer (5’-3’) (bp) AT1G20620 AGCTTCCAGTCAATGCTCCC ACCTGTCTCCAGCCTGTTTG 219

AT1G20620 CGACAAGCTGCTCCAGTGTA ACCTGTCTCCAGCCTGTTTG 476

AT1G20620 CGGTGTCCACACCTACACTC TTGGCCTCTAGATGCTTGGC 872

AT3G16640 CCAGGCTCAGTGACTTCCAA TTGGCCTCTAGATGCTTGGC 249 GGCAGGTAATTTTAATAGGG- AT3G16640 GCTGAAGAAGGTGGTGAGGAT 640 GACG

101 Table 3.2. Primer sequences used for PCR validation of expression in M. incognita egg, J2, and adult female cDNA libraries.

contig Amplicon Forward primer (5’-3’) Reverse primer (5’-3’) ID (bp) 1 CAGCCGGCTCTCTCGTTATT GTTTATCGCCAACAGCAGGC 230 23327 GTCTCGTCGGTCTTACCAGC TGAGGTCGCTACGGGTAATG 184 9110 CCAAGCCTGTGCTTGCTTTT CCAAGCCTGTGCTTGCTTTT 155 144 TGATGGGCGCAAAATGCAAA AGGAGAAGACCGGTTCTGGA 260 171 CCGCTGGCGGTGTTAAAGTA ACGTGCGTCTTGCTAAACCT 252 3702 GGCCCCGGAAATGATGATGA TTCCGATTCTCCAGGAGCCA 126 761 GATCCCGATGCGAAAAAGCC ACAGAGCCCCAATCTTCACG 212 7645 GATCCCGATGCGAAAAAGCC ACAGAGCCCCAATCTTCACG 212 1465 GGCCAGAATCTTGTCTGCCT GATCAATCGCACAACCTCCG 138 10276 AGCTTCCCGGCTTTCCATTT CCTTCCGAAGTTCGTGCTGA 255 22581 TCCGGGTGGACTGTATTGAC TGGTGGGCAGAACTTGTGAA 186 7898 CGCAAAGCAAATCCAAAAGCC GGAATCGAACTGGACTGCCA 238 1392 ACGTTGTAGGAGCAGGGTTG GACTACGTCGACGGAATGGT 278 14470 CGTGCTGGTTCCTTACTTGC CTCGGTAAGTTGGCGGAGTT 253 13050 GAGGCTCAAAACAACGCTGG TGAGAACCTGGCCGGAAAAG 197 16243 ATTGCTGCAAGCAGTTACCAC GGTGGAGGTGCAAGGAATGG 130 12435 TAGCCCCCGAGCTAACATCA CCCTCTTCGTGTCAGCAACA 161

102 64% no significant BLASTx hit

(17,906 contigs) Caenorhabditis (1,439 contigs)

Loa loa (1,355)

Brugia malayi (1,162)

Strongyloides (1,158)

plant-parasitic

other

2% Meloidogyne (332 contigs) (492 contigs) Heterodera (94) Bursaphenlenchus xylophilus (25) Globodera rostochiensis (23) Radopholus similus (10) Ditylenchus (9) Aphelenchoides (4) (1)

Figure 3.1. Graphical depiction of the distribution of top tBLASTx hits of the transcriptome of the Meloidogyne incognita adult female on Arabidopsis. The majority of contigs returned no significant BLASTx hit (E value < 10-3), illustrated in the top pie graph. The lower pie graph illustrates the plant-parasitic nematode top BLASTx hits, representing 2% of the transcriptome assembly.

103 # contigs 0 10 20 30 40 50 60 70 80 90 100

metabolic process ion transmembrane transport oxidation-reduction process G-protein coupled receptor signaling pathway protein folding proteolysis signal transduction transport

transmembrane transport single-organism process cation transmembrane transport cell redox homeostasis embryo development ending in birth or egg… neurological system process proteolysis involved in cellular protein… regulation of membrane potential

GO terms (Biological Process) (Biological terms GO single-organism cellular process synaptic transmission, cholinergic protein peptidyl-prolyl isomerization neurotransmitter transport neuropeptide signaling pathway determination of adult lifespan putatively secreted single-organism transport whole transcriptome cellular process ion transport

Figure 3.2. Top twenty common gene ontology terms (biological process) of putatively secreted proteins (blue) in adult female M. incognita transcriptome. Abundance of transcripts with these GO terms present in the whole transcriptome (red).

104 # contigs 0 20 40 60 80 100 120

integral component of membrane 294 membrane 224 extracellular space synapse plasma membrane endoplasmic reticulum

cell junction integral component of plasma… neuron projection endoplasmic reticulum membrane extracellular region cell membrane part acetylcholine-gated channel complex lysosome cytoplasm intracellular

GO terms (CellularComparment) dendrite membrane spliceosomal complex putatively secreted cell part whole transcriptome neuronal cell body membrane nucleus neuronal cell body anchored component of plasma…

Figure 3.3. Top twenty common gene ontology terms (cellular compartment) of putatively secreted proteins (blue) in female M. incognita transcriptome. Abundance of transcripts with these GO terms present in the whole transcriptome (red).

105 # contigs 0 20 40 60 80 100 120 hydrolase activity extracellular ligand-gated ion channel… catalytic activity calcium ion binding G-protein coupled receptor activity transferase activity, transferring glycosyl…

binding kinase activity page proceedingacetylcholine-activated the illustration. cation-selective… The figure/table number must appear on both the ½ title transferase activity page and the page with the illustration. hydrolase activity, hydrolyzing O-glycosyl… zinc ion binding acetylcholine binding metal ion binding carboxypeptidase activity acetylcholine receptor activity GO terms (Molecular Function) (Molecular GO terms oxidoreductase activity nucleotide binding peptidyl-prolyl cis-trans isomerase activity protein disulfide isomerase activity putatively secreted peptidase activity whole transcriptome cysteine-type endopeptidase activity

Figure 3.4. Top twenty common gene ontology terms (molecular function) of putatively secreted proteins (blue) in adult female M. incognita transcriptome. Abundance of transcripts with these GO terms present in the whole transcriptome (red).

106 Table 3.3. Top 15 most abundant transcripts expressed in adult M. incognita based on average coverage and paired-end reads mapped. Coverage/ Top BLASTx hit accession / Gene Ontology InterPro Contig E value Paired Reads Potential Function description Term Scan Mapped

AF273737/ Heterodera glycines 275,460/ 493 esophageal gland cell secretory 1.3e-31 - - effector 367,304 protein (hsp10)

228,786/ 10354 - - - - ? 283,426

M83820/ Xenopus laevis mucin 122,891/ 1223 3.3e-27 - cuticle B.1 229,896

107 473 114,322/ - - - - ? 532,994 JX863901/ Meloidogyne javanica IPR021109 92,729/ 24019 fatty acid and retinol-binding 1.6e-43 lipid metabolism IPR033121 195,960 protein IPR015818 LM036485/ Toxocara canis GO:0006869 87,193/ 13 5.9e-08 IPR015255 ? genome GO:0005319 859,440 IPR015819 79,343/ 386 - - - - ? 339,450

126 - - - - 68,119/ ? 420,484 61 - - GO:0006869 IPR001747 66,119/ lipid transport / egg yolk protein

Continued

107 Table 3.3 continued

Coverage/ Gene Top BLASTx hit accession / InterPro Paired Reads Contig E value Ontology Potential Function description Scan Mapped Term

58,567/ 564 - - - - ? 176,102 AF273737/ Heterodera glycines 55,303/ 3085 esophageal gland cell secretory 8.8e-20 - - effector 65,994 protein (hsp10) U35449/ tipulae GO:0006869 IPR001747 51,017/ *1733 9.9e-15 egg yolk protein vitellogenin (vit-6) gene GO:0005319 IPR011030 55,066

108 8 KRH17806/ Glycine max 41,516/ 7.01e-73 - - plant mimic hypothetical protein 1,846,844

39,468/ 181 - - - - ? 281,018 CAA65507/ Teladorsagia IPR002486 38,680/ 12 9e-26 GO:0042302 cuticle circumcincta cuticular collagen IPR008160 4475,448 - no significant BLASTx hit, no associated GO terms, no IPS match * contig has predicted signal peptide GO:0006869 (biological process): lipid transport GO:0005319 (molecular function): lipid transporter activity GO:0042302(molecular function): structural constituent of cuticle IPR021109: aspartic peptidase IPR033121: peptidase family A1 IPR001747: lipid transport protein IPR011030 / IPR015818 / IPR015255: vitellinogen IPR015819: lipid transport protein IPR002486: nematode cuticle collagen IPR008160: collagen triple helix repeat

108

Table 3.4. Top 15 most abundant transcripts in adult M. incognita with putative secretion signal, based on average coverage and paired-end reads mapped. Gene InterProSca Contig Top BLASTx hit accession / description E value Potential Function Ontology n endopeptidase 927 XM001983605/ Drosophila grimshawi 8.34e-04 GO:0004867 IPR002223 inhibitor XP013291202/ Necator americanus 828 0.0 - - ? protein disulfide-isomerase IPR002223 endopeptidase 183 XM002046423/ Drosophila virilis 3.68e-05 GO:0004867 IPR020901 inhibitor IPR000668 GO:0006508 IPR000169 683 GU360972/ cathepsin B 1.36e-75 digestion GO:0008234 IPR025660 IPR025661

109 IPR015819 249 KKA71696/ Pristionchus pacificus GO:0006869

1e-122 IPR015816 egg yolk protein vit-6, partial GO:0005319 IPR001846 XM003099104/ Caenorhabditis remanei 378 1.61e-11 - - ? hypothetical protein GO:0006030 1321 LN609530/ Strongyloides ratti 1.80e-14 GO:0008061 IPR002557 egg shell GO:0005576 XM002139351/ Cryptosporidium muris calcium- 987 3.28e-04 GO:0005509 IPR011992 signaling dependent kinase Continued 109 Table 3.4 continued

Gene Contig Top BLASTx hit accession / description E value InterProScan Potential Function Ontology 2676 XM003110555/ Caenorhabditis remanei 1.82e-13 - - ?

* AY714229/ Meloidogyne incognita GO:0006508 1 0.0 IPR001314 digestion chymotrypsin-like serine proteinase GO:0004252 XM003110555/ Caenorhabditis remanei 691 2.94e-14 - - ? hypothetical protein AJ557572/ Meloidogyne incognita putative 6920 6.7e-34 - - digestion cathepsin L protease XM002139351/ Cryptosporidium muris calcium- 1014 5.59e-05 GO:0005509 IPR011992 signaling dependent protein kinase 87 LM434076/ Nippostrongylus brasiliensis 3.18e-11 - - ? *144 ABC88426/ Meloidogyne incognita cathepsin D- GO:0006508 110 0.0 IPR033144 digestion like aspartic proteinase GO:0004190

* expression confirmed with RT-PCR - no associated GO terms, no IPS match GO:0005509: calcium ion binding IPR020901: proteinase inhibitor, Kunitz GO:0004252: serine-type endopeptidase GO:0004867: serine-type endopeptidase inhibitor IPR011992: EF-hand domain IPR015819: lipid transport protein IPR001314: chymotrypsin GO:0006508/GO:0006508: proteolysis IPR015816: vitellinogen IPR002557: chitin binding IPR001846: von Willebrand factor GO:0008234: cysteine-type peptidase IPR002223: pancreatic trypsin inhibitor Kunitz IPR033144: cathepsin D IPR025661: cysteine peptidase, histidine active site GO:0006869: lipid transport IPR000668: peptidase C1A GO:0005319: lipid transporter activity IPR000169: peptidase C1A, papain C-terminal GO:0006030: chitin metabolic process IPR025660: cysteine peptidase, cysteine active site GO:0008061: chitin binding GO:0005576: extracellular region

110 M. hapla C. elegans

D. vivaparus T. circumcincta

Figure 3.5. Pairwise comparison results (E value < 10-10) of RKN (M. incognita) 1,696 putatively secreted contig set with EST or cDNA databases from M. hapla, C. elegans, T. circumcincta, and D. viviparous.

111 Table 3.5. RT-PCR validation of expression detected in RNA-Seq dataset from adult female M. incognita. RT-PCR relative expression results from egg and J2 cDNA libraries included for life stage comparisons.

Functional Contig top tBLASTx hit Egg J2 Adult Category ID

16243 pectate lyase (M. enterolobii) + + ++ cell wall modifying 12435 endoglucanase (eng-2; M. incognita) + + + 1465 expansin b1 (M. incognita) - - +

1 chymotrypsin (M. incognita) + - + proteolytic 144 cathepsin-D (M. incognita) + + +

calcium- 761 §calreticulin (M. incognita) + low + binding

isomerase 7898 chorismate mutase (M. arenaria) + ++ ++

yolk protein 1392 vitellogenin-6 (Ascaris suum) - - ++

lipid acyl 14470 patatin (Trichinella spiralis) + + + hydrolase

diverse 23327 14-3-3b (M. incognita) low ++ + functions known 171 transthyretin-like (R. similis) + + +

22581 VAP-1 (G. rostochiensis) + + low 9110 VAP-2 (M. incognita) + + + function(s) 13050 amphid-secreted (G. rostochiensis) + + + unknown 7645 gland protein 23 (M. incognita) + low + 3702 gland protein 21 (M. incognita) - - low § Previously characterized virulence factor - Transcript not detected low = Transcript appears less abundant relative to controls + Transcript detected ++ Transcript appears more abundant relative to controls

112 A 1 2 3 B 1 2 3 C 1 2 3 D 1 2 3

E 1 2 3 F 1 2 3 G 1 2 3 H 1 2 3

Figure 3.6. RT-PCR analysis of M. incognita cDNA template derived from eggs (1), J2 (2), and adult females (3). β-actin used for reference (A), pectate lyase (B), Cre-vit-6 (C), patatin (D), CM (E), VAP-1 (F), amphid-secreted (G), and eng-2 (H) (100bp ladder; NEB).

113 A 1 2 3 B 1 2 3

1 2 3 1 2 3 C D E 1 2 3 F 1 2 3

Figure 3.7. RT-PCR analysis of M. incognita cDNA template derived from eggs (1), pre- parasitic J2 (2) and adult females (3). β-actin used for reference (A), chymotrypsin (B), 14-3-3b (C), VAP-2 (D), cathepsin-D (E) and expansin (F) (100bp ladder; NEB).

114 Table 3.6. Cell wall-degrading or modifying top BLASTx hits in female M. incognita transcriptome. Paired Reads Contig Top BLASTx hit accession / description E value Mapped/ Coverage

-39 1465 HQ386233.1|H. avenae expansin B1 2.45E 2876/437

-31 4180 AF127915.1|G. rostochiensis pectate lyase 1 precursor 3.41E 172/21

4421 HQ386233.1|H. avenae expansin B1 4.65E-37 1,544/259

4422 HQ386233.1|H. avenae expansin B1 9.67E-37 1,714/263

9649 HM798586.1|H. glycines expansin 1.54E-52 190/31

-24 9736 HQ915028.1|M. javanica expansin B 2.39E 486 /101

AF323086.1|M. incognita beta-1,4-endoglucanase -99 12435 1.51E 326/29 (eng-2)

-10 14734 EU143763.1|B. xylophilus expansin 1.42E 316/102

-14 15344 HQ915028.1|M. javanica expansin B 7.69E 216/42

-37 15688 FJ839965.1|H. avenae beta-1,4-endoglucanase (eng-1a) 4.10E 30/7

16243 HQ180169.1|M. enterolobii pectate lyase 1.22E-27 486/69

18432 AF527788.1|M. incognita pectate lyase 9.19E-31 14/7

18811 JQ768418.1|A. fragariae cellulase (eng-3) 3.61E-40 118/18

18899 HQ386233.1|H. avenae expansin B1 1.19E-36 118/28

19445 EU475875.1|M. incognita xylanase 2 5.51E-12 94/42

19927 EU475875.1|M. incognita xylanase 2 5.17E-18 94/29

21010 HQ915028.1|M. javanica expansin B 7.04E-19 78/17

22001 HQ915028.1|M. javanica expansin B 3.05E-99 364/65

Continued

115 Table 3.6 continued

Paired Reads Contig Top BLASTx hit accession / description E value Mapped/ Coverage 1.19E- 22002 HQ915028.1|M.e javanica expansin B 111 142/25

AF323086.1|M. incognita beta-1,4-endoglucanase 22306 8.00E-27 88/20 (eng-2)

AF323086.1|M. incognita beta-1,4-endoglucanase -27 22426 1.34E 54/15 (eng-2)

22622 HQ915028.1|M. javanica expansin B 3.98E-27 50/10

22644 EU143763.1|B.xylophilus expansin 3.49E-22 92/20

22898 AF323098.1|M. arenaria beta-1,4-endoglucanase (eng-2) 4.24E-06 72/18

HQ132749.1|R. reniformis GHF5 beta-1,4-endoglucanase -20 23226 5.75E 56/18 (eng-1)

23807 AY098646.1|M. incognita polygalacturonase 1.51E-82 40/12

2.59E- 24144 AF323086.1|M. incognita beta-1,4-endoglucanase (eng-2) 112/19 128

24220 EU475875.1|M. incognita xylanase 2 2.55E-12 16/8

24825 HQ915028.1|M. javanica expansin B 1.37E-18 102/21

25012 AF127915.1|G.rostochiensis pectate lyase 1 3.75E-37 202/26

AF049139.1|M. incognita cellulose binding protein 25194 1.63E-69 28/7 precursor cbp-1

25936 EU475875.1|M. incognita xylanase 2 4.92E-04 48/15

26213 EU475875.1|M. incognita xylanase 2 1.13E-07 78/38

26551 EU190885.1|R. similis xylanase 5.76E-12 40/21

26724 HQ123259.1|H. glycines pectate lyase (pel-7) 2.26E-09 24/7

26732 HM798586.1|H. glycines expansin 3.12E-44 62/22 Continued 116 Table 3.6 continued

Paired Reads Contig Top BLASTx hit accession / description E value Mapped/ Coverage

28554 AY098646.1|M. incognita polygalacturonase 6.05E-62 36/16

28574 AF323086.1|M. incognita beta-1,4-endoglucanase 2.04E-42 10/6

28649 HM798586.1|H. glycines expansin (expb1-1) 2.11E-18 20/10

30565 EU475875.1|M. incognita xylanase 2 6.26E-14 94/24

117 A 1 2 3 B 1 2 3 C 1 2 3

D 1 2 3 E 1 2 3

Figure 3.8. RT-PCR analysis of M. incognita cDNA template derived from eggs (1), pre- parasitic J2 (2) and adult females (3).β-actin used for reference (A), transthyretin (B), gland protein 21 (C), calreticulin (D), and gland protein 23 (E) (100bp ladder; NEB).

118 Table 3.7. Expression (+) of 19 genes in adult females of 27 pioneer genes identified in a gland cell-specific M. incognita cDNA library (Huang et al., 2003). (-) indicates the absence of this accession in our dataset and therefore has no associated E-value (N/A). Accession RNA-Seq E value / Number dataset tBLASTx bit score AY142120 - N/A AY142121 + 7.67e-16 / 92 AY142116 - N/A AF531161 + 1.91e-96 / 258 AF531164 + 8.99e-56 / 176 AF531165 + 2.71e-48 / 199 AY134432 + 2.15e-46 / 155 AY134442 + 9.08e-67 / 260 AY134437 + 3.83e-66 / 246 AF531160 + 1.33e-42 / 179 AY134433 + 6.66e-153 / 433 AY134431 - N/A AF531166 + 2.67e-61 / 242 AY134434 + 1.62e-08 / 37 AY134438 - N/A AY142119 + 1.13e-41 / 163 AF531167 - N/A AF531168 + 0 / 391 AF531169 + 3.05e-129 / 208 AY134435 - N/A AY134436 + 5.17e-136 / 193 AY134439 + 5.39e-118 / 263 AY134441 + 3.55e-33 / 77 AY134443 + 4.44e-70 / 270 AY134444 + 1.48e-73 / 283 AY135363 - N/A AY142118 - N/A

119 Table 3.8. Arabidopsis transcripts detected across four replicates of RNA-Seq analysis of M. incognita female cDNA libraries.

Unique Reads Gene ID Gene Description Mapped (Average)

AT5G60390 tu elongation factor 1607

AT3G41768 18s rRNA 227

AT5G09810 actin 7 18

AT3G53750 actin 3 15

AT4G05320 polyubiquitin 10 11

ATMG00020 mitochondrial 26S protein 8

AT2G42100 actin-like protein 8 translationally controlled *§ AT3G16640 5 tumor protein (TCTP) *AT1G20620 catalase 3 6

AT5G02500 heat shock protein 4

AT1G64740 alpha-tubulin 3

glyceraldehyde-3-phosphate AT1G13440 2 dehydrogenase

*expression verified by RT-PCR in at least two of three cDNA libraries § detected in only three of four RNA-Seq replicates

120 1 2 3 4 5 1 2 3 4 5

Figure 3.9. RT-PCR results of select A. thaliana transcripts from female M. incognita cDNA libraries. Lanes 1-3: catalase (AT1G20620; 219bp, 476bp, and 872bp). Lane 4-5:

TCTP (A T3G16640; 249bp and 640bp). Two independent replications in panels above, note negative result in second panel, lane 3. A transcript was considered “detected” when amplified in at least two independent cDNA libraries (100bp ladder; NEB).

121

CHAPTER 4: THE FEMALE SOYBEAN CYST NEMATODE SECRETOME & TRANSCRIPTIONAL RESPONSES TO PI 88788 RESISTANCE

4.1. ABSTRACT

Soybean cyst nematodes (SCN; Heterodera glycines) are the most yield-reducing pathogens of soybean in North America. Resistant soybean cultivars are widely utilized for management purposes; however, SCN populations are quickly adapting to overused sources of resistance. Counties across Ohio were surveyed for SCN and HG Type assays were performed to determine populations’ virulence to available sources of resistance. Of the populations tested to date, 96% are virulent on PI88788, which is the most prevalent source of resistance in commercial varieties. The objective of this study was to determine the differences in gene expression associated with SCN populations that have overcome this source of resistance. We hypothesize that transcriptional changes play a role in the adaptation to resistance, such that the expression of factors detected by the host are reduced, and factors that counteract or interfere with the host defense response are increased. Nine SCN populations from across Ohio were selected, which vary in their HG Type and level of virulence. These populations were grown on both PI88788 and a susceptible cultivar, Lee 74. Female SCN were harvested 15 days after inoculation with juvenile worms, total RNA was extracted, and submitted for RNA-Seq. Interesting expression patterns were found common among virulent populations such as putative effectors and genes related to digestion, the immune response, and detoxification. Also, populations avirulent on PI88788 share the expression of an effector almost entirely absent in virulent populations, indicating that the loss of this effector’s expression may play a role in virulence to this resistance source. Further work includes cloning of the

122 identified effector in order to test whether it hinders the infectivity of an otherwise virulent SCN population on PI 88788.

4.2. INTRODUCTION

Soybean cyst nematode (SCN; Heterodera glycines Ichinohe) is the most destructive pathogen of soybean (Glycine max (L.) Merr.), causing billions of dollars in yield loss in the United States alone (Wrather and Koenning, 2006). Supporting their sedentary endoparasitic niche, SCN initiate the formation of a complex feeding site referred to as a syncytium. As many as 200 root cells comprise one syncytial cell via the dissolution of cell walls and subsequent fusion of their cytoplasm (Jung and Wyss, 1999; Goverse et al., 2000). A diverse range and abundance of proteins are secreted by SCN into the host and essentially redirect plant pathways for the purposes of the feeding nematode (reviewed in Davis et al., 2008 and Gheysen and Mitchum, 2008). The use of host resistance is a favored management strategy in terms of both cost and sustainability, and has served this agricultural crop well. The introduction of the resistant cultivar Forrest, for example, was estimated to save the soybean industry upwards of $400 million in prevented yield losses (Bradley and Duffy, 1982 cited in Miller, 1986). Complete resistance to SCN is not available in soybean, making it imperative that sources of resistance currently available are used wisely in order to prolong their effectiveness. In order to make wiser management decisions with sources of host resistance, a classification scheme was devised to describe SCN populations’ ability to reproduce on resistant lines. SCN populations are classified by an “HG Type” test (HG for Heterodera glycines) based on their ability to grow to maturity on available sources of resistance originating from seven different plant introduction lines (Niblack et al., 2002). Virulence in this parasite is defined by their relative reproductive ability on resistant versus susceptible lines; the resulting percentage is referred to as the population’s Female Index (FI). When the FI is greater than 10%, the population is considered virulent to that resistant line as denoted by its HG Type. For instance, if a population has a FI > 10% on

123 indicator lines 2, 5, and 7 (PI 88788, PI 209332, and PI 548316, repectively) it is classified as HG Type 2.5.7. Despite the identification of virulent SCN shortly after the introduction of resistant soybeans (Ross and Brim, 1957), host resistance has remained an enormously valuable tool for the management of the soybean cyst nematode. However, the genetic variability of SCN has allowed populations to quickly adapt to overused sources of resistance (Dong and Opperman, 1997; Niblack et al., 2002; Niblack et al., 2008). The genetic bases for two major sources of resistance to SCN have been recently deciphered. Underlying Plant Introduction (PI) 88788-derived resistance is an allele of the quantitative trait locus, Rhg1 (resistance to Heterodera glycines), rhg1-b. The three genes that underlie this locus are a putative amino acid transporter (Glyma18g02580), an α-SNAP protein (Glyma18g02590), and a protein with a WI12 domain (wound-inducible protein 12), which lacks any functionally characterized domains (Glyma18g02610). These three genes are present in susceptible varieties; however, the presence of 10 tandem copies of the set of genes confers the resistance phenotype (Cook et al., 2012). Another important quantitative trait locus, Rhg4, is conferred by a serine hydroxymethyltransferase, wherein two polymorphisms affect the enzyme’s activity in the susceptible versus resistant phenotype (Liu et al., 2012). The current understanding is that Rhg4 resistance requires Rhg1 for full function, for reasons not yet understood (Meksem et al., 2001). The most prevalent sources of SCN resistance in commercially available varieties are derived from PI 88788 and Peking (reviewed in Concibido et al., 2004). These two resistance sources vary in their action against SCN specific to the nematode’s feeding site. Peking-based resistance affects the earliest stages of feeding site initiation, resulting in the death of the developing infective second-stage juvenile (J2) to the J3 stage. PI 88788-based resistance in comparison, is a more prolonged reaction, affecting the later J3 and J4 stages (Klink et al., 2010). J2 SCN penetrate soybean lines with rhg1-b, however the initiation or maintenance of their feeding sites fail, which ultimately prevents many of them from maturing into reproductive adults (Li et al., 2004). The examination of later stages of infection may help to illuminate the genetic mechanisms that allow SCN

124 populations to successfully overcome this resistance. Specifically, changes in nematode gene expression during late infection may be an important component of adaptations to PI 88788-derived resistance. Transcriptional studies of SCN have focused predominantly on the nematode’s esophageal gland cells, which are known to secrete many of the proteins involved in parasitism. Early morphological observations indicated that the subventral glands were more active during initial stages of parasitism, with the dorsal gland becoming the predominant source of secretions during the later stages of parasitism (Bird, 1983; Hussey and Mims, 1990). Chemical stimulation of the metacorpus, subtractive hybridization, and direct microaspiration of the gland cell contents have been used to isolate and analyze gene expression in these cells (Jaubert et al., 2002; Gao et al., 2001; Gao et al., 2004). Numerous parasitism transcripts were identified, including cell wall- modifying enzymes acquired via horizontal gene transfer, and plant peptide mimics (Bakhetia et al., 2007; Bekal et al., 2003; Wang et al., 2005; Wang et al., 2010). Also numerous in these studies are protein-encoding genes with no significant homology to known proteins or domains. These transcripts have been referred to as ‘pioneers.’ Most of their functions remain to be elucidated but are presumably somehow involved in this highly intricate parasitic interaction. In order to successfully parasitize a host, SCN must evade both innate and induced plant defense responses directed at the feeding site as well as the nematode itself (Smant and Jones, 2011). Because penetration and migration do not seem to be hindered during nematode infection of PI 88788, maintenance of the feeding site is likely more relevant to virulence during this particular resistance response. In order to identify nematode genes involved in parasitism on different resistant lines, genetic analysis of SCN has been made possible through controlled crosses of inbred lines in (Dong and Opperman, 1997). For example, two alleles of the chorismate mutase effector were identified, wherein Hg-cm-1A (or a closely linked gene) is selected for when nematodes are grown on PI88788, but not on the cultivar Hartwig, PI90763, or the susceptible cultivar Lee 74 (Bekal et al., 2003; Lambert et al., 2005). Two more genes involved in parasitism were recently implicated in SCN virulence on PI 88788, a SNARE-like protein

125 (HgSPL-1, KM575849) and a biotin synthase (HgBioB) (Bekal et al., 2015). Bekal et al. (2015) verified expression of HgSPL-1 in the subventral esophageal glands and also implicated its potential interaction with the Rhg1-encoded α-SNAP protein through co- expression and purification studies. Researchers also observed a reduced copy number of HgSPL-1 in virulent populations. This evidence suggests that HgSPL-1 may function as an avirulence protein. As depicted in Jones and Dangl’s (2006) ‘zigzag’ model of the plant immune system, pathogens avoid effector-triggered immunity by either discarding or diversifying the effectors that have been recognized by the host. Examples of these strategies have been documented in bacterial, fungal, and oomycete pathogens (Kousik and Ritchie, 1996; Liu et al., 2013; Pais et al., 2013). Due to the obligately sexual reproduction of SCN, the field populations isolated for this study represent multiple genotypes, thus limiting our ability to make meaningful sequence comparisons, but analysis of differential gene expression between populations could detect other adaptations, such as the disappearance of an effector. One hypothesis of how changes in gene expression may contribute to overcoming host resistance is that genes are upregulated to effectively counteract the resistance response. This could be observed in an abundance of transcripts encoding detoxification enzymes that would lessen the stress experienced in syncytia during the resistance response. The expression of proteins that inhibit plant defense pathways could also be expected. Another valid hypothesis predicts that the loss of expression of a gene responsible for host detection would also result in host defense evasion. The primary objective of this study was to determine the relative expression of genes in SCN populations virulent versus avirulent to PI 88788-resistance, grown on the susceptible cultivar Lee 74. Gene expression differences between virulent versus avirulent populations will be further analyzed for their potential roles in overcoming this source of resistance.

126

4.3. MATERIALS AND METHODS

4.3.A. Nematode population collection and maintenance Nematode populations were collected directly from the field in counties across Ohio and from field samples submitted to the C. Wayne Ellett Plant and Pest Diagnostic Clinic (The Ohio State University; Columbus, Ohio). Populations chosen for this study came from Brown, Darke, Defiance, Erie, Hancock, Henry, Knox, Lucas Madison, Pike, and Wood counties. Cysts were isolated from soil samples with a semi-automatic soil elutriator (University of Georgia Instrument Shop) over a 30-mesh (600 μm aperture) sieve atop an 80-mesh (180 μm aperture) sieve. Cysts were then ground with a rubber stopper against an 80-mesh sieve over a 500-mesh (25 μm aperture) sieve to retain the eggs, which were then separated from debris via sucrose flotation (Jenkins, 1964). Populations were maintained on the susceptible soybean cultivar, Lee 74, grown in sand- Turface (1:1 v/v) (Turface MVP; Buffalo Grove, IL) amended with a slow-release fertilizer. Prior to setting up an experiment, eggs were freshly extracted and used to infest a “start” pot of Lee 74 soybean seedlings. Eggs were harvested from start pots approximately thirty days after infestation, following the extraction procedure described above, to infest experimental plants which were placed in crock containers set in a water table maintained at 27°C (Fig. 4.1.A.).

4.3.B. HG Type assays PI 88788 and Lee 74 soybean seed were sterilized with chlorine gas in a vacuum chamber overnight. Seed were rolled in germination paper saturated with water. After three days, uniform germinated seedlings were chosen for planting. Eggs were extracted from a start pot as described in section 4.3.A. Four thousand eggs, constantly agitated on a stir plate for consistency in concentration, were used to infest each transplanted seedling. Assays were harvested 28 to 32 days after infestation, cysts were extracted as described in section 4.3.A., and sucrose flotation was used to separate them from debris. Cysts were counted under a dissecting microscope. Assay results were only recorded

127 when the average cyst count on the susceptible cultivar Lee 74 was ≥ 100. A Female Index was calculated to determine the population’s HG Type as described by Niblack et al. (2002). HG Type assays are routinely used for populations to ensure that virulent populations are maintained.

4.3.C. Female SCN isolation Eggs from each population were harvested from start pots as described in section 4.3.A. Eggs were placed on a 25 μm aperture nylon mesh, saturated with water and allowed to hatch. Five days after eggs were placed into the hatching chamber, J2 were collected from the water below the nylon mesh. Infective juveniles (1,000) of each population were used to infest 32 soybean seedlings in individual cone-tainers of PI 88788 and Lee 74. Cone-tainers were split between 32 individual crocks (16 planted with 10 cone-tainers of PI 88788 and 16 with 10 cone-tainers of Lee), and crocks were placed in a greenhouse water table (27°C) in a completely randomized design. Fifteen days after infestation, four samples consisting of four plants of an individual cultivar and population from four individual crocks were pooled for female extraction via elutriation and sucrose flotation (described in section 4.3.A). This time point was chosen in order to optimize the number of females isolated prior to melanization (Fig. 4.1.B.). Immediately following elutriation, females were kept on ice until RNA extractions were performed. Not all populations grown on PI 88788 produced a sufficient number of females for RNA isolation; these included Darke, Defiance, Hancock, and Madison.

4.3.C. RNA isolation and Illumina library synthesis Total RNA was extracted from three independent pools of 300 adult female soybean cyst nematodes (SCN) following the RNeasy Plant Mini Kit protocol (Qiagen, Valencia, CA). Homogenization of the tissue was performed with an eppendorf pestle in RTL buffer (10% β-mercapto-ethanol). RNA quantity and quality were assessed with a Qubit® 2.0 Fluorometer (Life Technologies, Carlsbad, CA) and a Bioanalyzer (Agilent Technologies, Palo Alto, CA), respectively. All samples selected for cDNA synthesis had RNA Integrity Numbers (RIN) values ≥ 7.

128 RNA from each pool (1.5 μg) was used to generate an adaptor-tagged double- stranded DNA library for RNA-Seq with the TruSeq Stranded mRNA Sample Prep LS protocol (Illumina, San Diego, CA) following the manufacturer’s protocol. Quantification of DNA was done with the Qubit® 2.0 Fluorometer (Life Technologies, Carlsbad, CA) and quality was assessed with the Bioanalyzer (Agilent Technologies, Palo Alto, CA). The pooled samples were diluted to 4 nM and combined to generate one multiplexed DNA library for each replicate.

4.3.D. Illumina sequencing and transcriptome assembly The cDNA libraries (10 pmoles) were sequenced on two flow cell lanes of the Illumina HiSeqTM2000 platform at the Ohio State University Comprehensive Cancer Center (OSUCC). Both ends of the library were sequenced to generate 100 nt raw paired- end reads. The Illumina Analysis Package CASAVA 1.8.2 was used to perform bcl conversion and demultiplexing. Image deconvolution and quality value calculations were carried out with the Illumina GA pipeline v1.8. The SCN raw reads were imported into CLC Genomics Workbench (v6.5.1, CLC Bio) and trimmed for quality, adapter indexes and poly(A) tails with the default settings (Ambiguous limit = 2, quality limit = 0.05). Processed reads were assembled de novo by means of two independent approaches. First, the CLC Bio algorithm based on de Bruijn graphs and the optimized parameters [Word Size = 54, Bubble Size = 650, Length Fraction = 0.6, Similarity Fraction = 0.95] was used to assemble contigs of ≥250 nt; and second was the Oases v0.2.08 (Schultz et al. 2012) with Kmer sizes of 53, 59, 65, 71, 77, 83, and 89. Transcripts with ≥90% sequence similarity were collapsed into clusters and the longest read retrieved with CD-HIT-EST (Li and Godzik, 2006). The assemblies were then merged into a final assembly using Minimus2 (Sommer et al., 2007). The raw sequence reads will be submitted to the NCBI short sequence read archive. Preprocessed reads were aligned to the reference gene sets with the map to reference function in CLC bio Genomics Workbench with the following parameters: Similarity Fraction = 0.95; Length Fraction = 0.95; default settings herein. Genes differentially expressed between populations and hosts were identified with DESeq2

129 (Anders and Huber, 2010), which determines changes in transcript accumulation based on a negative binomial distribution model. Significance was defined at an FDR < 0.05 (Benjamini and Hochberg, 1995) for all analyses unless otherwise specified.

4.3.E. SCN secretome analyses The six possible open reading frame (ORF) amino acid sequences were predicted from the transcript sequences of SCN with ORF-Predictor (Min et al., 2005). Only the subset of predicted sequences ≥50 amino acids was used in subsequent analyses. The SignalP 4.1 neural networks algorithm (Nordahl Peterson et al., 2011) was implemented to detect putative transmembrane proteins with signal peptide secretion and cleavage site signatures in their amino acid sequences according to the default settings for D-score. De novo assembled transcripts containing a putative secretion signal were assigned hierarchical gene ontologies (GO terms) on the basis of biological processes, molecular functions, and cellular components via the platform-independent Java™ 6 implementation of the BLAST2GO software (Gotz et al., 2008). The top five tBLASTx hits to the nr database with a cut-off E-value of 10-3 were considered for GO annotation. BLAST2GO software was also used to run InterProScan software (Mitchell et al., 2015) to determine the presence of functional domains and Enzyme Code mapping for all putatively secreted proteins.

4.3.F. Reverse transcriptase PCR for expression validation. RNA isolation for RT-PCR experiments was done in the same manner as described in section 4.3.C. RNA template (500ng) was treated with RQ1 DNase (Promega, Madison, WI) and used for cDNA synthesis with the GoScriptTM Reverse Transcription System (Promega, Madison, WI) following the manufacturer’s protocol, with an oligo(dT)15 primer. During cDNA synthesis, a no-RT reaction was included for each RNA sample for detection of contaminating genomic DNA. cDNA template was diluted 2:1 and amplified with Phusion® High Fidelity DNA Polymerase (New England Biolabs, Ipswitch, MA). Oligonucleotides were synthesized by Integrated DNA Technologies (Coralville, IA, U.S.A.); primer sequences used are listed in Table 4.1. The

130 PCR cycle conditions used were: initial denaturation at 98°C for 30 seconds; 30 cycles of 98°C for 30 seconds, 57°C for 15 seconds, and 72°C for 10 seconds; and a final extension at 72°C for 10 minutes.

4.4. RESULTS

4.4.A. SCN population virulence Nine different soybean cyst nematode populations collected from across the state of Ohio (namely, Brown, Darke, Defiance, Erie, Hancock, Knox, Madison, Pike, and Wood County) were selected based on the results of HG Type assays (Fig 4.2). Those chosen for this study represent HG Type 7, 5.7, 2.5.7, 1.2.5.7 and 1.2.3.5.7 and also differ from one another in their Female Index (Fig. 4.3).

4.4.B. Sequence assembly Each sample represents nematodes harvested from four plants pooled (a total of 42 samples were run on two flow cell lanes). The pooled cDNA libraries were sequenced, which generated over 1.14 billion paired-end, 100 nt reads. After trimming (quality, adapters, poly(A) sequences), ~827.3 million reads were obtained (694.4 million reads in pairs). The de novo assembled transcriptome consisted of 38,118 non-redundant SCN transcripts (≥226 nt; x̅ = 899 nt).

4.4.C. The female SCN secretome Transcript sequences were translated into their putative amino acid sequences. A total of 31,327 SCN sequences of ≥50 amino acids were identified. In silico analysis of transcripts coding for secreted proteins with SignalP (Nordahl et al., 2011) revealed 1,843 (5.9%) SCN sequences with signatures for signal peptide secretion and cleavage sites. Additionally, 11% of the putatively secreted proteins had a transmembrane domain signature.

131 For the subset of putatively secreted proteins, 870 (47%) had significant BLASTx matches to the nr database. Functional categorization of this set of genes resulted in 106 gene ontology (GO) terms describing 771 contigs in the sub-ontology, “cellular compartment.” The top two GO terms, ‘integral component of the membrane’ and ‘membrane’ represented 65% of the contigs classified in this sub-ontology (Fig. 4.4). The top 20 terms for the sub-ontology “biological process” represented approximately 40% of this set of genes, with the most abundant terms predictably being associated with membrane systems and transport (Fig. 4.5) as this stage represents a feeding female. Just over 40% of genes with associated GO terms in the sub-ontology “molecular function,” are represented by the top 20 terms, which includes functions indicative of the parasitic interaction and digestion such as transmembrane transport, and processes related to metabolism and oxidation-reduction (Fig. 4.6). With BLAST2GO software (Gotz et al., 2008), top BLASTx hits against the non- redundant database were determined for the putatively secreted proteins. Roughly 60% of the proteins returned significant BLASTx hits with an E-value cut-off of 10-3. For the translated transcripts with significant hits to the nr database, homology to Toxocara canis, Ascaris suum, Strongyloides ratti represented nearly 40% of the contigs annotated (Figure 4.7). Hits to Heterodera glycines accessions were the twelfth most abundant, likely reflecting the disproportionate representation of this plant-parasite in the non- redundant (nr) database in relation to the animal-parasitic nematodes.

4.4.D. The female SCN transcriptome DESeq2 analysis resulted in the detection of 4,340 differentially expressed genes (Anders and Huber, 2010). Among these, almost 40% (1,636 contigs) of the expression patterns were shared between two or more of the populations (Figure 4.8). Only 1.5% (67 contigs) of the differentially expressed genes were shared between all of the populations grown on the susceptible versus resistant soybean lines. An even smaller portion of these transcripts, representing only 0.002% (9 contigs), appeared to be commonly regulated between the avirulent population (Hancock) versus all of the virulent populations.

132 Transcripts found to be upregulated in the avirulent population in comparison to all of the virulent populations includes a protein with a CYTH-like domain, a group which is known to act on triphosphorylated substrates, and an ATP-binding cassette (ABC) transporter (Table 4.2). Other transcripts present in higher abundance in the avirulent population were a putative zinc finger nuclease, a murein transglycosylase with a tetraspanin domain, and a transcript containing immunoglobin and fibronectin domains (Table 4.2). Most intriguingly is the presence of transcripts with homology to a dorsal gland protein (H. avenae, ADI82807.1), which is nearly absent from all the virulent populations sequenced. The presence of the transcript in this population was verified in another replication of the experiment with reverse transcriptase PCR (Fig. 4.9). The transcript was successfully amplified in the avirulent population Hancock (HG Type 7) as well an additional avirulent population, Henry (HG Type 5.7), but absent in Lucas (HG Type 7). The amplicon was only faintly detected in the virulent population, Defiance (HG Type 2.5.7) and absent in the other virulent population tested, Brown (HG Type 1.2.5.7). Transcripts in greater abundance in all of the virulent populations relative to the avirulent population (Hancock) were also detected (Table 4.3). This included a putative ferritin with GO terms associated with iron binding and transport. Two additional transcripts had no significant BLASTx hit, but one did return InterPro scan results of a SNARE coiled-coil domain and plectin repeat (IPR000727) with gene ontologies associated with protein binding and the cytoskeleton (GO:0005515, GO:0005856). Bekal et al. (2015) recently identified a protein of similar identity and produced evidence implicating it as an as an avirulence factor. Alignment of the translated sequence (432 amino acids) with that of the SNARE-like protein (KM575849) identified by Bekal et al. (2015) shows homology to the N-terminal, first 160 of 326 amino acids with 34% identity (Fig. 4.10). Other genes, uniquely downregulated in HG Type 2.5.7 populations relative to the avirulent population include a gag-pol type transposon and another contig with a top BLASTx hit to the gland specific protein (ADI82807.1) (Table 4.4); genes upregulated include a cathepsin B-like protease, two twichin-like genes, and another containing a fibronectin domain (Table 4.5).

133 Due to the differences in resistance reactions observed in Peking-type resistance and PI 88788-type resistance (Klink et al., 2010; Li et al., 2004), we also examined the differential expression unique to the HG Type 7 population versus HG Type 1.2.5.7 and 1.2.3.5.7 populations. Among interesting genes downregulated in these virulent populations are retrotransposons and an annexin domain-containing protein (Table 4.6). Uniquely upregulated genes in these virulent populations also include an annexin-like protein (Table 4.7). Patterns in gene expression were also analyzed for females of each population grown on the resistant line PI 88788 versus the susceptible line, Lee 74. The avirulent population, Hancock, was not included in this analysis due to insufficient female isolation from PI 88788. Aside from an abundance of hypothetical proteins, many transcripts in this set encode enzymes potentially involved in detoxification, such as peroxiredoxin, thioredoxin, glutathione S-transferase, and oxidoreductase (Table 4.8). There were only two down-regulated genes common to all populations grown on PI 88788 versus Lee 74 (or conversely, upregulated on Lee 74), one containing a zinc finger domain (IPR013083) and the other contig having no significant BLASTx hit or InterProScan identified domain.

4.5. DISCUSSION

The sources of resistance available against SCN primarily fall into two major categories. One category includes the indicator lines 1, 3, and 6 (PI 548402, PI 90763, and PI 89772) and to some extent, indicator line 4 (PI 437654) (Colgrove and Niblack, 2008). Among this first category, the resistance response observed in the cultivar Forrest (resistance from PI 548402) includes cell death surrounding the syncytium, disconnecting the feeding site, within which the cytoplasm condenses, and ultimately starvation of the feeding nematode. The other major category of SCN resistance resides in indicator lines 2, 5, and 7 (PI 88788, PI 209332 and PI 548316). In this category of resistance, such as that observed in the cultivar Bedford (PI 88788-derived), nuclear degeneration is

134 observed prior to condensing of the cytoplasm (Kim et al., 1987). Another noteworthy difference between these categories of resistance is the stage in which an effect is observed on the invading nematode. Resistance in the 1, 3, 6-category hinders the earlier second and third stage juveniles whereas resistance in the 2, 5, 7-category predominantly affects the development of third and fourth stage juveniles (Halbrendt et al., 1992). Thus, some adaptations to PI 88788 resistance potentially express themselves during the later stage of nematode development. Studies focusing on soybean cyst nematode’s secretome have contributed tremendously to our understanding of this parasitic interaction. The majority of parasitism-related proteins are secreted from the nematode’s esophageal gland cells and delivered through the stylet. This focus has been fruitful because the interaction predominantly relies upon the interface between the nematode’s stylet and the plant cell’s plasma membrane, where its feeding site is formed. However, for this sedentary endoparasitic nematode, the interface of the interaction likely extends beyond its head region, as its entire body is surrounded by host tissue. Other important secretory tissues are the hypodermis, which deposits substances onto the surface of the cuticle, and amphids, which are most notable for their role in chemosensing during migration. Secreted cuticular proteins have been identified in other cyst nematodes as important factors in parasitism (Jones et al., 2000; Robertson et al., 2000). Amphids were discovered to secrete an avirulence protein by another sedentary endoparasite, the root-knot nematode (Semblat et al., 2001). Phasmids are similar in structure to amphids but located posteriorly, and are also sources of secretions (reviewed in Perry 1996). A homologue of a C. elegans putatively phasmid-secreted protein was identified in infective juvenile root-knot nematode secretions (Bellafiore et al., 2008). Rectal glands are another likely source of secretions that are important during the parasitic interaction, as they are presumably involved in root-knot nematode parasitism during egg laying (Rosso et al., 1999). Many studies that focus on secretions from the esophageal glands have isolated their contents via microaspiration or even isolated intact gland cells for transcriptomic analysis (Gao et al., 2001; Maier et al., 2013). Others have chemically induced pharyngeal pumping to analyze the proteome of those secretions (Bellafiore et al., 2008).

135 The former methods are problematic at later life stages when the living female cyst remains partially embedded in host tissue, and the latter method is impossible due to this stage’s obligate nature. The whole female nematode transcriptomic approach used in our study allowed us to capture the expression of genes not only in esophageal glands but also in the other important secretory tissues mentioned above. This approach also allowed us to examine an under-studied life stage, which provides us with new candidate genes involved in parasitism and clues to a potential mechanism involved in adaptations to PI 88788 resistance. Our discovery of a dorsal gland effector-like protein expressed in avirulent populations and nearly absent in virulent populations suggests that one factor populations may have used to adapt to PI 88788 resistance is to suppress the expression of an effector in order to evade detection and the elicitation of host defenses. Bekal et al. (2015) decidedly did not pursue a similar transcript, with 62 amino acids of a 162 amino acid fragment homologous to Ha-dsl-1 (AD182806.1) since the same allelic form was present in both virulent and avirulent inbred populations. This sequence remains unpublished but is presumably unique from our effector-like protein (AD182807.1), but a sequence comparison could be informative. The presence of the effector amplicon in the virulent population “Defiance” may be explained by the fact that this adaptation is an example of the loss or reduction of expression rather than the loss of a gene or functional protein. Alternatively, a portion of the Defiance population may contain individuals not yet adapted to this resistance source. The down-regulation or complete loss of effectors has been documented in other pathosystems (reviewed in Cui et al., 2015). It will be interesting to investigate whether there is any detectable fitness cost to loss of this effector’s expression, in addition to whether its presence indeed hinders a virulent population’s parasitic success on PI 88788. Other adaptations to resistance have been discovered through genetic analyses, such as the allele of a chorismate mutase effector, Hg-cm-1A that is selected for when nematodes are exclusively grown on PI88788 (Lambert et al., 2005). Our experimental design does not allow for a genetic analysis between the populations isolated around Ohio, as they represent multiple genotypes. But genes with differential expression

136 between virulent and avirulent populations would be good candidates for genetic analyses to determine the mechanism underlying these patterns of expression. Another significant limitation to our study is the inclusion of only one avirulent population. This likely inflates the number of differentially expressed genes shared in those comparisons. Ongoing work is underway to repeat this RNA-Seq experiment with the inclusion of additional avirulent populations. In addition to the upregulation of an effector-like gene, the avirulent population presents other upregulated genes of interest. Fibronectin in the cuticle of root-knot nematode has been shown to potentially play a role in Pasteuria penetrans endospore attachment through hydrophobic interactions (Mohan et al., 2001; Davies et al., 1995). Twitchin is a polypeptide located in muscle cells of C. elegans that includes fibronectin and immunoglobin domains, which are also known to play a role in microbe interactions (Benian et al., 1993). A couple twitchin-like transcripts were observed in the set of genes upregulated in HG Type 2.5.7 populations compared with the avirulent population. While containing fibronectin domains, these twitchin complexes are more likely involved in the regulation of muscle contraction, not with organismal interactions based on work done in C. elegans (Benian et al., 1993). Among interesting transcripts in greater abundance in virulent populations is an uncharacterized protein (contig_4434) with a SNARE coiled-coil domain (IPR000727) and protein binding and cytoskeleton gene ontologies (GO:0005515, GO:0005856). Alignment of the translated sequence (432 amino acids) with the SNARE-like protein (KM575849) identified by Bekal et al. (2015) that has been implicated as an avirulence factor, aligns to the first (N-terminal) 160 of 326 amino acids at 34% identity (SFig.4.2). The uniqueness of the protein in our dataset may provide some role in virulence that has not yet been detected by the host. As proposed by Bekal et al. (2015), diversity in these SNARE–like proteins may be a result of the interaction with the resistance related α- SNAP protein in Rhg1. Aside from an abundance of hypothetical proteins, many transcripts upregulated on PI 88788 encode enzymes potentially involved in the elevated detoxification that may be required to endure the resistance response, such as peroxiredoxin, thioredoxin,

137 glutathione S-transferase, and oxidoreductase. Similarly, a Meloidogyne incognita- secreted glutathione S-transferase has been implicated in the detoxification of phytoalexins in Arabidopsis (Dubreuil et al., 2007). There were only two down-regulated genes common to all populations grown on PI 88788 versus Lee 74, one containing a zinc finger domain (IPR013083) and the other contig having no significant BLASTx hit or InterPro-identified domain identified. These genes may be actively repressed during the resistant response or alternatively may be induced in the interaction with the susceptible Lee 74. In addition to the nematode response to resistance, a transcriptional profile of this adult female stage may also provide valuable insight into other intersecting aspects of this parasite’s biology. In H. glycines, the adult female body becomes a protective case, shielding the eggs within from adverse environmental conditions (or simply lack of a suitable host) for years in the field. The mature female’s dead body, full of eggs, is what is referred to as the “cyst.” The processes that occur relating to the melanization of the female body upon death are not well understood. Mechanisms of cuticular melanization discovered in insects may provide some direction into categories of proteins that are likely to be involved. For instance the phenoloxidase laccase-2 was determined to be essential to cuticle tanning in the red flour beetle (Arakane et al., 2005). While no laccase-2-like top BLASTx hits are observed in our transcriptome, three contigs were found to share significant homology with the pea aphid laccase-1. The possibility of SCN sharing a similar phenoloxidase, essential to the melanization of the maturing cyst should not be discounted. Our data suggests that gene expression associated with avirulent versus virulent SCN populations may play a role in the adaptation to PI 88788 resistance. Ongoing work is underway to repeat this RNA-Seq experiment with the inclusion of more avirulent populations, which will likely narrow our lists of differential gene expression common to avirulent versus virulent populations. Future work on candidate genes should include cloning and expression of SCN-secreted proteins in planta to begin investigating their role in this parasitic interaction. Also of interest is to further explore other clues pertinent to this parasite’s biology. The origin and localization of secreted proteins from adult

138 female SCN could also be investigated as it relates to other important aspects of parasite biology such as cuticle melanization.

4.6. ACKNOWLEDGMENTS We acknowledge Dr. Bryan Cassone for the bioinformatic analysis of RNA-Seq data. This work was supported in part by funding from The United Soybean Board, Ohio Soybean Council, and The Ohio State University, Department of Plant Pathology and Center for Applied Plant Sciences, SoyRes Research Team.

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144 3 dpi 15 dpi

A) B) 18dpi 24 dpi

Figure 4.1. 32 soybean seedlings in individual cone-tainers were infested with infective juveniles of each soybean cyst nematode population in crocks containing either PI 88788 or susceptible Lee 74 (A). Three days post infestation, acid fuchsin-stained juveniles can be observed (white arrow). The time point, 15 days post infestation (dpi), was chosen to optimize the number of females isolated prior to obvious melanization (18 and 24dpi).

145 Table 4.1. Primer sequences used for detection of transcripts in female soybean cyst nematode cDNA libraries.

Gene ID Forward primer (5’-3’) Reverse primer (5’-3’)

CGCTGAACCCGAAGG- TTGATGTCACGGACGA-

β-actin (AF318603) CCAACAGA TCTCACG putative DG-secreted AAAGTGGTGGCACGGA- TTGGGCGATGACCATTTGGA (contig_19995) GAAA

146 Hg Type 7 Hg Type 5.7 Hg Type 2.5.7 Hg Type 1.2.5.7 Hg Type 1.2.3.5.7

Figure 4.2. Soybean cyst nematode populations across Ohio were selected based on variation in geographic location, virulence, and HG Type. Populations include both HG Type 2 indicated in yellow, green, and blue, and non-HG Type 2 indicated in red and orange.

147 300

250

200 FI = 40

150 FI = 30 Brown Pike # Cysts PI on 88788 Cysts # 100 Knox FI = 20 Defiance Wood Madison

50 FI = 10

Darke Lucas Hancock 0 Henry 0 100 200 300 400 # Cysts on Lee

Figure 4.3. Soybean cyst nematode populations vary in virulence to both the susceptible and resistant host; means from one HG Type assay are displayed. Color corresponds to population’s HG Type, red = HG Type 7, orange = 5.7, yellow = 2.5.7, green = 1.2.5.7, and blue = 1.2.3.5.7. Female Index (FI) thresholds are indicated with dotted lines (assays resulting in < 100 cysts on the susceptible checks are considered invalid).

148 # of contigs 0 100 200 300 400 integral component of membrane membrane

collagen trimer Golgi membrane endoplasmic reticulum integral component of plasma membrane nucleus intracellular gap junction endoplasmic reticulum membrane cytoplasm plasma membrane mitochondrion mitochondrial inner membrane Golgi apparatus ribosome Gene ontology term (Cellular Compartment) (Cellular term ontology Gene extracellular space extracellular region intracellular membrane-bounded organelle oligosaccharyltransferase complex cytosol signal peptidase complex trans-Golgi network nucleosome proton-transporting V-type ATPase, V0 domain

Figure 4.4. Top twenty common gene ontology terms (cellular compartment) of putatively secreted proteins in female soybean cyst nematode transcriptome.

149 # of contigs 0 10 20 30 40 50 60 70 80 90 transmembrane transport metabolic process oxidation-reduction process

transport ion transport nematode larval development proteolysis translation embryo development ending in birth or egg… signal transduction locomotion protein phosphorylation carbohydrate metabolic process protein glycosylation phosphorylation G-protein coupled receptor signaling pathway

Gene ontology term (Biological Process) (Biological term Gene ontology multicellular organism development neurotransmitter transport lipid metabolic process determination of adult lifespan

Figure 4.5. Top twenty common gene ontology terms (biological process) of putatively secreted proteins in female soybean cyst nematode transcriptome.

150 # of contigs 0 5 10 15 20 25 30 35 40 45 50 protein binding ATP binding structural constituent of cuticle zinc ion binding

metal ion binding transporter activity catalytic activity nucleotide binding structural constituent of ribosome ATPase activity, coupled to transmembrane… calcium ion binding oxidoreductase activity transferase activity, transferring acyl groups hydrolase activity transferase activity transmembrane transporter activity binding

Gene ontology term (Molecular Function) (Molecular term Gene ontology hedgehog receptor activity nucleic acid binding heme binding neurotransmitter:sodium symporter activity transferase activity, transferring glycosyl groups

Figure 4.6. Top twenty common gene ontology terms (molecular function) of putatively secreted proteins in female soybean cyst nematode transcriptome.

151 no significant hit Toxocara canis (196 contigs) Ascaris suum (131) Strongyloides ratti (122 38% (702 contigs) Caenorhabditis (102) Ancylostoma ceylanicum (73) Brugia malayi (68) Loa loa (61) Haemonchus contortus (60) plant-parasitic other (270)

3% Heterodera glycines (28 contigs) (58 contigs) Meloidogyne (11)

Globodera pallida (9)

Heterodera avenae (3)

Bursaphelenchus xylophilus (3)

Ditylenchus destructor (2)

Globodera rostochiensis (1)

Radopholus similis (1)

Figure 4.7. Species distribution of top BLASTx hits of SCN putatively secreted contigs in female soybean cyst nematode transcriptome.

152 1,636

2,704

DEG 4,340 Unique

non-DEG ≥ 2 Populations 13,232 Resistant vs. Susceptible Avirulent vs. Virulent

Figure 4.8. Approximately ¼ of the female soybean cyst nematode transcriptome represents differentially expressed genes (DEG) between populations grown on a susceptible versus resistant line. Of the DEG, approximately 60% are unique to one population and the remaining DEGs are common to ≥2 populations. Sixty-seven DEGs are shared between all populations grown on the susceptible versus resistant line (green). Nine DEGs are common between the avirulent and virulent populations (orange).

153 Table 4.2. Differentially expressed genes downregulated in populations virulent to PI 88788 relative to the avirulent soybean cyst nematode population, “Hancock,” grown on the susceptible cultivar, Lee 74.

Fold Top BLASTx Hit InterPro Gene Ontology p value Contig # Change Description Scan Term (adj.)* (Log2)*

19995 dorsal gland protein no IPS none predicted -3.30 2.52E-05

PR012337 GO:0003676 13888 zinc finger protein -2.49 6.25E-04 IPR008906 GO:0046983 CYTH-like domain 5148 IPR027417 none predicted -4.76 2.24E-12 containing

41597 ABC transporter no IPS none predicted -5.20 2.41E-04

murein 32087 IPR018499 GO:0016021 -4.48 4.94E-06 transglycosylase IPR013783 34003 no significant hit GO:0005515 -4.56 1.08E-06 IPR003961

* fold change (log2) and p value (adjusted) are based on averages of comparisons IPR012337: ribonuclease H-like domain IPR008906: HAT, C-terminal dimerization domain IPR027417: p-loop containing nucleoside triphosphate hydrolase IPR018499: tetraspanin/peripherin IPR003961: fibronectin type III IPR013783: immunoglobulin-like fold GO:0003676: nucleic acid binding GO:0046983: protein dimerization activity GO:0016021: integral component of membrane GO:0005515: protein binding

154 1 2 3 4 5 1 2 3 4 5 NTC

A B

Figure 4.9. A) Amplification of putative dorsal gland-secreted effector observed in lane 4, faint amplicon observed in lanes 3 and 5. B) Amplification of β-actin in corresponding samples for reference. 1: Lucas (HG Type 7), 2: Brown (HG Type 1.2.5.7), 3: Defiance (HG Type 2.5.7), 4: Hancock (HG Type 7), and 5: Henry (HG Type 5.7) (100bp ladder, NEB; NTC = PCR no template control).

155 Table 4.3. Differentially expressed genes upregulated in soybean cyst nematode populations virulent to PI 88788 relative to the avirulent SCN population, “Hancock,” grown on the susceptible cultivar, Lee 74.

Top BLASTx Hit InterPro Gene Ontology Fold Change p value Contig # Description Scan Term (Log2) (adj.) GO:0006826 *8083 putative ferritin IPR009078 GO:0006879 1.95 1.06E-03 GO:0008199 IPR000727 GO:0005515 4434 no significant hit 1.82 3.50E-12 IPR001101 GO:0005856

*187 no significant hit no IPS none predicted 1.41 6.12E-04

*contig contains predicted signal peptide IPR009078: ferritin-like superfamily IPR000727: target SNARE coiled-coil homology domain IPR001101: plectin repeat GO:0006826: iron ion transport GO:0006879: cellular iron ion homeostasis GO:0008199: ferric iron binding GO:0005515: protein binding GO:0005856: cytoskeleton

156 Query 1 MAPKCLPLELLFEIVPFIPAEKAAPNALSSCLLLHNLLLPRVIKW-KELKKMIKELRDEV 59 M PK LP ELLFE+V FIP E A PN SSC LLHN+L RV+KW KE+ + +LR E Subject 1 MTPKFLPPELLFELVLFIPVEDAIPNVFSSCWLLHNILHSRVVKWEKEMVNFMNKLRGEN 60 Query 60 FGKIDELRDEMNQKFGQINTRLDRTDQRLDQMDKRLDRM-DQRLDQFEQYGPVPPPMPLQ 118 I +LR E + I D+ + ++D+ D++ DQ + +Q + Subject 61 DQTIKQLRAENAELRSVIAELKDQVANQYVRIDELKDQVADQNVRIHDQKKEIRKLREDN 120 Query 119 SGHYSGIESTSEF-----SELRQIGTSAAFDTRQNTGSELQRRPPF 159 + + + ST + R+I ++ A Q+ GSE + PF Subject 121 QKNATFMRSTKGLLCEIKGQNRRINSAMAMFREQHFGSETCQEHPF 166

Figure 4.10. Alignment of translated contig_4434 (subject) and the SNARE-like protein identified by Bekal et al. (2015), KM575849 (query). Figure adapted from blasp suite- 2sequences run on the NCBI webserver.

157 Table 4.4. Genes uniquely downregulated in the virulent soybean cyst nematode populations (HG Type 2.5.7; Darke, Defiance, Erie, and Madison) relative to the avirulent population (HG Type 7; Hancock).

Fold p value Contig Top BLASTx Hit Description Accession E Value Change (adj.) (Log2)

1520 calcium-independent phospholipase ERG79792.1 9E-40 -2.52 1.33E-08

24762 Transposon Ty3-I Gag-Pol polyprotein KRY45026.1 2E-09 -3.00 4.75E-03

21313 dorsal gland protein ADI82807.1 4E-41 -3.15 1.34E-04

33582 uncharacterized protein XP_007243180 1E-15 -4.75 6.20E-10

12 contigs with no significant BLASTx hits not displayed above

158 Table 4.5. Genes uniquely upregulated in the virulent soybean cyst nematode populations (HG Type 2.5.7; Darke, Defiance, Erie, and Madison) relative to the avirulent population (HG Type 7; Hancock). Fold p value Contig # Top BLASTx Hit Description Accession E Value Change (adj.) (Log2) 773 cathepsin B-like proteinase AIG62903 5e-07 1.18 2.50E-01

12105 carbohydrate kinase WP_027459835 4e-29 1.62 5.10E-02 leucine-rich repeat-containing 22326 ERG79687 6e-14 protein 1.03 2.50E-01 108 uncharacterized protein XP_014097857 6e-04 1.80 2.70E-01 4922 XP_016849353 RNA helicase 7e-11 2.31 2.91E-01 9377 fibronectin type III domain protein XP_013300813 2e-138 1.86 7.11E-05 5667 Twitchin KHN73749 5e-58 1.88 1.40E-12 5322 Twitchin KHN73749 3e-52 1.45 1.44E-03 5322 Twitchin KHN73749 3e-52 1.45 1.44E-03 24 contigs with no significant BLASTx hits not displayed above

159 Table 4.6. Genes uniquely downregulated in the virulent soybean cyst nematode populations HG Type 1.2.5.7 (Brown and Wood) and 1.2.3.5.7 (Knox and Pike)* relative to the avirulent population, HG Type 7 (Hancock).

Top BLASTx Hit Fold Change p value Contig # Accession E Value Description (Log2) (adj.)

10990 ran-binding protein 4 AFD54151 2e-55 -1.32 4.54E-04

Pao retrotransposon 12095 XP_001898012 0.0 -1.07 2.21E-04 peptidase DNA polymerase epsilon 12429 KOO32988 3e-09 -2.90 2.57E-03 catalytic subunit zinc finger BED domain- 13131 XP_016340910 1e-15 -1.97 2.44E-03 containing protein

15462 G protein-coupled receptor KKA71288 2e-4 -4.05 2.44E-03

18032 dorsal gland protein ADI82807 1e-43 -1.16 2.55E-02

18196 dorsal gland protein ADI82807 3e-27 -5.12 5.65E-04

26522 adenylyl cyclase V CEF67186 8e-14 -2.32 4.78E-03

§29287 dorsal gland protein ADI82807 3e-29 -2.48 1.20E-03

4073 putative gland protein AAO85459 3e-27 -3.50 2.35E-10

peptidase M12A and 4399 CDJ82742 3e-15 -3.09 6.99E-08 AMMECR1 domain

6446 cation efflux family protein EJW79974 1e-98 -3.79 1.79E-04

annexin repeat domain 6479 CDJ94946 2e-37 -6.42 1.82E-19 containing

6456 hypothetical protein Hgg-29 AAL78223 8e-59 -1.04 2.16E-02

6949 ran-binding protein 4 AFD54151 1e-04 -3.26 1.24E-02 putative RNA-binding 7823 CDJ88382 3e-07 -3.83 2.81E-03 protein 8408 dorsal gland protein ADI82807 3e-13 -1.16 8.36E-08 Continued 160 Table 4.6 continued Top BLASTx Hit Fold Change p value Contig # Accession E Value Description (Log2) (adj.)

8821 serine proteinase CAA74206 4e-50 -3.06 8.62E-04

8883 glutathione synthetase XP_003142623 4e-69 -1.32 3.62E-03

9408 replication factor c complex ERG83470 2e-15 -1.01 2.24E-02

zinc finger BED domain- 9552 XP_016656274 2e-14 -3.76 1.44E-02 containing

§9771 putative cuticular collagen CAB88203 1.85e-49 -1.35 2.23E-02

RNA-directed DNA *30927 polymerase with integrase CDJ93006.1 0.0 -4.53 5.63E-04 domain

*34696 unnamed protein product CBY13755 7e-05 -3.73 7.86E-03

*uniquely downregulated genes in HG Type 1.2.3.5.7 populations relative to the HG Type 7 population §contig contains putative signal peptide

161 Table 4.7. Genes uniquely upregulated in virulent soybean cyst nematode populations (HG Type 1.2.5.7; Brown and Wood)* relative to the avirulent population (HG Type 7; Hancock). Fold p value Contig # Top BLASTx Hit Description Accession E Value Change (adj.) (Log2) 13032 ran-binding protein ADW77537 8e-61 3.91 2.97E-05

16912 annexin B9 XP_974030 3e-39 2.06 1.21E-03

4618 carnitine o-palmitoyltransferase ERG82347 1e-104 2.30 3.46E-27

17968 zinc carboxypeptidase-like protein KPM08353 2e-17 1.83 4.86E-03

34390 DREV methyltransferase KJH41147 1e-13 2.15 2.08E-02

12230 vitellogenin-6 ERG79133 2e-06 1.28 1.42E-02

6368 general substrate transporter CDJ80856 2e-52 1.46 1.12E-02

21189 general substrate transporter CDJ80856 2e-52 1.50 1.10E-02

24584 copine family protein XP_001891884 1e-29 1.48 2.55E-02 pleckstrin homology domain- 11261 XP_012154981 9e-94 1.09 5.43E-03 containing family *no uniquely upregulated genes were identified in HG Type 1.2.3.5.7 (Knox and Pike).

162 Table 4.8. Differentially expressed genes, upregulated in all soybean cyst nematode populations grown on the resistant versus susceptible cultivar. Results do not include the HG Type 7 population (Hancock), as not enough females were produced for RNA isolation. BLASTx hit Contig ID Top BLASTx hit description E value Accession contig_806 hypothetical protein Tcan_05511 [T. canis] 1E-17 KHN84509.1 contig_2777 hypothetical protein Tcan_05510 [T. canis] 2E-26 KHN84508.1 contig_915 hypothetical protein CBG01305 [C. briggsae] 4E-19 XP_002636065.1 contig_4269 hypothetical protein Hgg-29 [H. glycines] 9E-62 AAL78223.1 contig_9059 serine proteinase [H. glycines] 1E-81 CAA74206.1 contig_5248 2E-18 contig_5558 unknown [H. glycines] 1E-16 AAW33665.1 contig_4745 2E-18 contig_66 unknown [H. glycines] 1E-06 AAW33667.1 contig_3755 tricarboxylate carrier [N. americanus] 2E-134 ETN82230.1 contig_9 C-type lectin domain protein [H. glycines] 0E+00 AAM18623.1 contig_4066 protein ADT-2, isoform c [C. elegans] 0E+00 NP_001024534.2 contig_3592 aspartic protease 2B [S. ratti] 7E-43 ACR56786.1 contig_390 peroxiredoxin [M. incognita] 2E-107 ACZ67203.1 contig_23 thioredoxin and Erv1 Alr domain [H. contortus] 4E-128 CDJ87420.1 contig_1335 lysine histidine transporter 1 [A. suum] 0E+00 ERG84129.1 contig_22142 26S protease regulatory subunit 4 [T. canis] 1E-01 KHN89100.1 4-hydroxyphenylpyruvate dioxygenase contig_7994 0E+00 ETN80809.1 [N. americanus] contig_6423 protein B0403.6 [C. elegans] 3E-90 NP_001024323.1 contig_1791 hypothetical protein CRE_14746 [C. remanei] 3E-02 XP_003101175.1 contig_2322 nose resistant to fluoxetine protein 6 [T. canis] 7E-78 KHN82264.1 contig_7038 0E+00 prolyl 4-hydroxylase alpha-related [S. ratti] CEF59606.1 contig_5692 5E-170 contig_1577 solute carrier family 28 member 3 [T. canis] 0E+00 KHN88807.1 saccharopine dehydrogenase-like oxidoreductase contig_332 2E-174 ERG79103.1 [A. suum] contig_2937 serine carboxypeptidases [R. similis] 7E-152 AIC75882.1 contig_2583 phospholipase b-like 2 [A. suum] 0E+00 ERG82527.1 contig_219 gut esterase 1 [A. suum] 7E-117 ERG83158.1 contig_5593 alpha-N-acetylgalactosaminidase [T. canis] 0E+00 KHN82558.1 contig_1162 protein ALH-4, isoform a [C. elegans] 3E-169 NP_741553.1 contig_152 chitin binding domain-containing [S. ratti] 8E-14 CEF70986.1 contig_841 protein CBR-TYR-5 [C. briggsae] 9E-125 CAP33379.2 contig_2117 protein Bm8880, isoform b [B. malayi] 1E-10 CDQ04478.1 Continued 163 Table 4.8 continued BLASTx hit Contig ID Top BLASTx hit description E value Accession contig_5848 Na(+)/H(+) exchange regulatory cofactor 1E-79 KHN71588.1 contig_94 putative phospholipase B-like 2 [S. ratti] 0E+00 CEF65923.1 contig_5484 C-1-tetrahydrofolate synthase [T. canis] 0E+00 KHN83783.1 contig_396 transthyretin-like family-containing [S. ratti] 5E-34 CEF59337.1 contig_6781 CRE-SCAV-3 protein [C. remanei] 8E-94 XP_003097233.1 contig_1243 collagen alpha-1(XX) chain [T. canis] 8E-96 KHN85683.1 contig_2 vitellogenin-6 [A. suum] 3E-146 ERG79133.1 contig_11927 hypothetical protein LOAG_15914 [Loa loa] 1E-09 EFO12619.2 contig_1980 glutathione synthetase [T. canis] 4E-100 KHN87278.1 contig_2056 lysine histidine transporter-like 4 [A. suum] 1E-140 ERG87556.1 contig_3723 protein DHS-2, isoform c [C. elegans] 4E-82 NP_871815.1 contig_1176 CBR-TYR-1 protein [C. briggsae] 1E-86 XP_002642685.1 contig_423 acyl-CoA synthetase family member 3 [S. ratti] 2E-168 CEF70664.1 contig_1168 ShTK domain containing protein [B. malayi] 4E-74 XP_001893282.1 contig_2030 hypothetical protein CRE_06721 [C. remanei] 1E-63 XP_003109874.1 contig_3061 sugar transporter SWEET1 [T. canis] 1E-67 KHN81692.1

164

CHAPTER 5: CHALLENGES, IMPLICATIONS, AND FUTURE DIRECTIONS

Symbiosis, in the most general sense, is a physically close and long-term interaction between organisms of different species. This interaction exists along a spectrum from the interdependency of mutualism to the often quite harmful parasitism. Obligate parasitism represents an extreme in symbiotic relationships, wherein one organism relies entirely on another for its own survival, to the detriment of that other individual. Obligate parasites navigate a tricky relationship with their host; in order for the infection to be successful the host must be alive. If a parasite exhausts its host’s resources, it may kill its host before it has accomplished its ultimate goal of producing progeny. Research on obligate parasites of agricultural plants in particular serves two major purposes: to protect and defend crops against these parasites and to broaden our understanding of the biology underlying the delicate balance struck between parasite and host. It is then our hope to take this understanding and apply it back to our first purpose, protecting our supply of food, fuel, and fiber. Root-knot and soybean cyst nematodes are the most economically damaging nematodes of crops around the globe. Root-knot nematodes (RKN; Meloidogyne spp.) are capable of parasitizing virtually every type of crop that is grown. While soybean cyst nematode (SCN; Heterodera glycines), as the name suggests, favors mainly the roots of soybean, its host range extends well beyond soybean. RKN and SCN are both difficult to manage once present in a field, but for different reasons related to their life histories and available management strategies. Crop rotation options are limited for RKN control due to their broad host range, and effective host resistance is only found in certain crops. Crop rotation and host resistance are the most effective strategies currently available for controlling SCN populations; however, cysts can maintain viable eggs in the soil for years, so rotation strategies are geared more towards managing populations rather than

165 attempting to eliminate them. Host resistance has also made a huge impact on controlling SCN populations, but the overuse of too few sources of resistance is pressuring populations to quickly overcome them. Chemical nematicides can be used to control both RKN and SCN, but this strategy is rarely cost effective and its use is becoming increasingly less common due to concerns over effects on the environment and human health. Genetic engineering is exploring multiple opportunities to develop novel methods of control. One strategy is to express small RNAs that are complementary to a nematode transcript, targeting it for degradation by RNA interference (RNAi) pathways.

Suppression of host RNAi during nematode parasitism

I examined RNAi pathways during the root-knot nematode-plant interaction and found evidence that the host’s RNAi pathways are being suppressed specifically in the feeding site during infection and that this suppression impacts both major pathways of RNAi, short interfering and micro-RNAs. I have also collected evidence that the suppression of RNAi benefits nematode parasitism, as the expression of some proteins known to suppress RNAi render plants more susceptible to infection. It is possible that nematodes directly interfere with these pathways with a secreted protein, a strategy that is employed in other plant-pathogen systems, including virtually all plant viruses, but also some bacterial pathogens and an oomycete. Perhaps equally likely is that the suppression of RNAi is an indirect effect of the reprogrammed root cells that make up the nematode feeding site. Both giant cells and syncytia exhibit qualities of developmental reprogramming which is now understood to be under the direct regulation of RNAi. Thus the physiological changes induced in nematode feeding sites on one hand could necessitate the suppression of RNAi or on the other it could be causing its suppression. The challenge to determining how this suppression is enacted is two-fold. Our understanding of RNAi pathways, while growing tremendously in the era of next-generation sequencing capabilities, is still in its infancy. And our understanding in model systems such as Arabidopsis may not translate to other systems. When analyzing

166 tobacco expressing the viral suppressor, HC-Pro, I observed an increase in susceptibility to RKN parasitism. Analyzing a similar component expressed in Arabidopsis did not appear to have an effect on the interaction. Preliminary results of a silenced reporter gene in this system have also shown no indications of feeding site-specific interference. Future work may benefit from examining the expression of other viral suppressors in Arabidopsis, driven by nematode-induced or feeding site-specific promoters. Directing the viral suppressor’s expression to the time or location of infection may decrease other off-target disturbances to the whole plant, which may otherwise indirectly affect the interaction. The other challenge lies in the great unknown of uncharacterized proteins secreted by nematodes. Both RKN and SCN have large repertoires of secreted proteins that remain uncharacterized. No homology to characterized viral suppressors was evident in our transcriptome datasets; however, this is not terribly surprising considering the diversity of known suppressors in both sequence and mode of action. RNAi suppressor proteins have now been characterized across kingdoms, from viruses to bacteria, and most recently from an oomycete. Taking a similar approach to plant virologists, effectors could be cloned and tested in vitro for suppressor capabilities. This type of screen should start with putative nematode effectors, which are expressed and secreted, including those that have previously been characterized, as suppression of RNAi could be an additional function. My findings implicating the suppression of RNAi by plant-parasitic nematodes also raise the interesting possibility that virus-vectoring nematodes may actually benefit from this seemingly self-serving association. Nematode-viral associations are known to be non-essential for nematode parasitism, but could they be beneficial? Whether nematodes in the order Dorylaimida suppress RNAi pathways during their parasitic interaction should be investigated. These relationships could also be explored for positive effects on nematode fitness initially by testing the susceptibility of hosts to nematodes vectoring viruses and viruses lacking a functional viral suppressor. RNAi is becoming an increasingly utilized tool for functional studies and is actively being explored as a control method for insects as well as nematodes. A better understanding of how RNAi is suppressed in plant-parasitic nematode feeding sites may

167 help researchers improve the efficacy of this technique. Many functional analyses have observed only a moderate decrease in target transcript abundance, which could be a result of insufficient processing in the nematode feeding site. If efficacy relies more on processing within the nematode, perhaps double-stranded or micro-RNA progenitors could be better targeted for nematode ingestion. Ingestion in these parasites however, is not well understood. While researchers have identified important effectors relevant during later stages of infection, a lot remains to be learned about the mature female nematode.

Transcriptomic analysis of female root-knot nematodes

Examining the transcriptome of the adult female root-knot nematode serves multiple purposes. Due to its endoparasitic lifehistory, this later stage has been less extensively studied relative to the parasitic juvenile and the responses in the surrounding host tissue. At this time point we expect to better observe gene expression related to the maintenance of giant cells, continued evasion and protection against host defenses, ingestion and digestion, and egg production. My transcriptomic analysis of the female root-knot nematode revealed the expression of putative genes involved in parasitism that may be relevant to this particular stage of parasitism. My study confirmed the expression of 19 of 27 previously identified in esophageal gland-specific cDNA libraries from multiple life stages. Expression observed in our dataset included genes encoding cell wall-modifying enzymes, various proteases, and previously characterized virulence factors. Other secreted proteases may be localized to the nematode intestine to aid in digestion; however, some proteases are known to be secreted into the host and inhibit defense pathways. The roles of protease- encoding genes in my dataset would need to be experimentally resolved. Determining the location of the transcript via in situ hybridization would provide good indication as to its role in digestion versus host interactions. The expression of the cell wall modifier, expansin, is of particular interest because it seems to be specific to this later female stage.

168 The role of cell wall modifying enzymes in later stages of infection has yet to be explored. Do cell wall modifying enzymes continue to play a significant role at this later stage of parasitism? How does surrounding root tissue accommodate the expansion of the female body? Is cell wall loosening required for later female development in the confines of the cortex and for the egg mass to access the external environment? In addition to discovering these interesting trends in gene expression, this transcriptomic analysis of female M. incognita has generated new candidate genes for functional analysis and has put forth new hypotheses to investigate. Genes of interest will be cloned so that their role in the parasitic interaction can be investigated. These results have also provided evidence that the feeding tube size exclusion limit should be revisited and that RNA analysis may be a suitable tool to begin this investigation. Detection and identification of molecules ingested by the female nematode is challenging due their limited abundance, making RNA an attractive molecule to search for because it can be amplified. While the vast majority of sequences in our dataset are of nematode origin, even transcripts of low abundance may be detected via PCR amplification. The presence of plant-derived transcripts of up to 872 nucleotides was found within the female nematode, just under 300kDa, far exceeding the currently accepted size exclusion limit. This finding highlights the need to readdress this perspective of nematode feeding. Does a transcript’s subcellular location affect its likelihood of uptake? The endomembrane system surrounding the feeding tube structure may make transcripts with endoplasmic reticulum retention signals of interest. New technologies improving our ability to follow messenger RNA molecules in vivo with targeted RNA binding proteins could be used to determine the subcellular localization of the transcripts identified in this study. Molecular weight is likely not a sufficient parameter with which to characterize the passage of molecules through the nematode feeding tube, as it is no indication of a substance’s shape or pertinent chemical properties. The expression of proteins tagged with a human influenza hemagluttinin epitope (HA) could be used to test how proteins’ different sizes and properties affect their entry. Future work examining the passage of

169 substances through the nematode feeding tube may ultimately help improve delivery mechanisms for nematicidal toxins or dsRNA molecules.

Transcriptomic analysis of female soybean cyst nematodes: differences in expression between virulent and avirulent populations

Researchers have recently deciphered the genes underlying the most commonly used resistance sources, PI 88788 and Peking. Populations of soybean cyst nematode continue to rapidly adapt to overused sources of resistance such as that derived from PI 88788. Over 96% of the populations tested from Ohio counties to date are classified as virulent to this indicator line. Results from my transcriptomic analyses of SCN populations across the state of Ohio suggest that differences in gene regulation may play a role in how SCN populations have adapted to PI 88788-based resistance. A finding of particular interest is the presence of a transcript with homology to a dorsal gland-specific protein in the avirulent population, but mostly absent from virulent populations. The adaptation to this resistance source may include the loss of expression of this effector, thus allowing evasion of detection and subsequent host defense responses. Future studies will benefit from the inclusion of more SCN populations, particularly those that are still avirulent to PI 88788 resistance, to verify the consistency of these findings. Is the dorsal gland-specific effector-like protein present in all avirulent populations tested? Cloning and expression of this effector in the PI 88788 soybean will help determine whether it hinders virulent populations’ ability to successfully reproduce on this line. Additionally, RNAi knockdowns of this nematode gene in avirulent populations could help determine whether its loss confers virulence to this type of resistance.

170 Future directions

In the age of next generation sequencing our ability to quickly identify candidate genes involved in parasitism has increased exponentially. Technologies for functional analysis have also improved considerably with the ability of RNAi strategies to knockdown gene expression in parasitic nematodes. Many questions remain regarding the late infection interactions of root-knot and soybean cyst nematodes with their hosts. What other nematode secretions are essential for the long-term maintenance of giant cells and syncytia? Do some secretions serve to allow the expanding female body and egg mass to emerge from the root? Are cellular contents taken up preferentially from feeding sites for ingestion? Answers to questions like these may allow researchers to develop more efficient tools for both functional analysis and novel or improved methods for nematode control.

171

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