PLANT-NEMATODE INTERACTIONS AND THE APPLICATION OF RNA
INTERFERENCE FOR CONTROLLING ROOT-KNOT NEMATODES
By
PHUONG THI YEN DINH
A dissertation submitted in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
WASHINGTON STATE UNIVERSITY Department of Plant Pathology
MAY 2015
© Copyright by PHUONG THI YEN DINH, 2015 All Rights Reserved
© Copyright by PHUONG THI YEN DINH, 2015 All Rights Reserved
To the Faculty of Washington State University:
The members of the Committee appointed to examine the dissertation of
PHUONG THI YEN DINH find it satisfactory and recommend that it be accepted.
______Debra A. Inglis, Ph.D., Chair
______Charles R. Brown, Ph.D.
______Lori M. Carris, Ph.D.
______Axel A. Elling, Ph.D.
______Kiwamu Tanaka, Ph.D.
ii
ACKNOWLEDGMENTS
I thank my parents, Nghia Dinh and Yen Nguyen, for their support, care and encouragement during my education. I especially want to express my deepest gratitude to Dr. Axel Elling, my major advisor for his incredible advice and persistent support. My sincere gratitude also goes to Dr. Debra Inglis, my committee member and my major advisor in the last year of my PhD program, for her encouragement, concern and guidance. I also am thankful to other committee members, Drs. Charles Brown, Lori
Carris, Kiwamu Tanaka and Brenda Schroeder, for their suggestions about my research and reviews of my manuscripts and dissertation. I extend my appreciation to Dr. Michael
Knoblauch for his guidance and support in microscopy techniques. I also express my gratitude to Professor Elizabeth Siler for her assistance with and proofreading of my dissertation.
I thank the faculty, staff, and students in the Department of Plant Pathology,
Franceschi Microscopy and Imaging Center as well as Plant Growth Facilities for their help, assistance, and friendship during my studies at Washington State University. I offer a warm thank-you to all members and undergraduate students, past and present, of Dr. Elling’s lab. It has been a great pleasure to work with these lab members. Thanks to all of my friends for their comfort. I am thankful to my colleagues in Southern
Horticultural Research Institute, Vietnam for their encouragement during my studies.
This project was funded by the Washington State Department of Agriculture,
Washington State Potato Commission, Idaho Potato Commission, Northwest Potato
Research Consortium, Washington Grain Commission, and United States Department of Agriculture, and I gratefully acknowledge the support.
iii
PLANT-NEMATODE INTERACTIONS AND THE APPLICATION OF RNA
INTERFERENCE FOR CONTROLLING ROOT-KNOT NEMATODES
Abstract
by Phuong Thi Yen Dinh, Ph.D. Washington State University May 2015
Chair: Debra A. Inglis
Plant parasitic nematodes are significant pests in major agricultural systems.
Sedentary endoparasites like root knot nematodes (RKN; Meloidogyne sp.) obtain plant- derived nutrients from feeding sites formed in host roots thereby supporting nematode development. The formation of feeding sites is hypothesized to be mediated by nematode effectors, proteinaceous secretions from nematode esophageal gland cells.
An inherent challenge in studying most plant-nematode interactions is the difficulty in directly observing nematodes within plant roots. Traditional microscopy techniques are unusable because developing nematodes are surrounded by layers of root cells. In this study, a novel, nondestructive technique was developed to observe the progression of nematode pathogenesis in planta. Pratylenchus penetrans, Heterodera schachtii and Meloidogyne chitwoodi were fluorescently labeled with the lipid specific stain PKH26 and inoculated onto Arabidopsis thaliana seedlings growing in microscopy rhizosphere chambers. The migration patterns and morphology of live nematodes then were observed using confocal microscopy during the parasitic life cycles.
Host-nematode interactions were studied at the molecular level by characterizing the highly conserved RKN effector, 16D10. Overexpression of 16D10 in
iv
A. thaliana increased the transcript level of VND7, a xylem development marker, and metaxylem root cell numbers, thereby enhancing susceptibility of A. thaliana to
M. incognita. Modifying xylem development by 16D10 possibly facilitates M. incognita feeding site formation. An ortholog of 16D10, Mc16D10L, was cloned from M. chitwoodi.
Plant-mediated 16D10 RNA interference (RNAi) silenced Mc16D10L and significantly reduced M. chitwoodi race 1 reproduction by up to 71% in A. thaliana and potato
(Solanum tuberosum cvs Russet Burbank and Désirée) plants. Introducing 16D10 RNAi into potato breeding line PA99N82-4 also decreased reproduction of M. chitwoodi pathotype Roza by 50%; this pathotype breaks RMc1(blb), a resistant gene of PA99N82-4.
The RNAi effect of Mc16D10L was transmitted to M. chitwoodi offspring, and significantly reduced pathogenicity of nematode offspring on non-RNAi plants. The potato RNAi line, D21 further proved resistant to M. incognita, M. javanica, M. arenaria and M. hapla. Plant-mediated 16D10 RNAi offers a promising new tool for molecular breeding against RKN in potato.
v
TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS ...... iii
ABSTRACT ...... iv
LIST OF TABLES ...... x
LIST OF FIGURES ...... xi
CHAPTER 1: GENERAL INTRODUCTION ...... 1
1. 1. Plant parasitic nematodes ...... 1
1. 1. 1. Root-lesion nematodes ...... 2
1. 1. 2. Cyst nematodes ...... 3
1. 1. 3. Root-knot nematodes ...... 6
1. 1. 4. Molecular plant-nematode interactions ...... 8
1. 1. 5. Management ...... 14
1. 2. Root-knot nematodes in potato ...... 23
1. 3. Research objectives and contributions ...... 25
1. 4. Literature cited ...... 30
CHAPTER 2: NONDESTRUCTIVE IMAGING OF PLANT-PARASITIC NEMATODE
DEVELOPMENT AND HOST RESPONSE TO NEMATODE PATHOGENESIS ...... 62
2. 1. Abstract ...... 63
2. 2. Introduction ...... 64
2. 3. Materials and methods ...... 66
2. 4. Results ...... 70
2. 5. Discussion ...... 74
vi
2. 6. Acknowledgments ...... 79
2. 7. Literature cited ...... 80
2. 8. Figures ...... 88
CHAPTER 3: MELOIDOGYNE INCOGNITA EFFECTOR 16D10 MODIFIES XYLEM
DIFFERENTIATION IN ARABIDOPSIS THALIANA TO FACILITATE NEMATODE
PARASITISM ...... 97
3. 1. Abstract ...... 98
3. 2. Introduction ...... 99
3. 3. Materials and methods ...... 102
3. 4. Results ...... 112
3. 5. Discussion ...... 117
3. 6. Acknowledgments ...... 123
3. 7. Literature cited ...... 124
3. 8. Figures ...... 134
3. 9. Supplement ...... 141
CHAPTER 4: RNA INTERFERENCE OF EFFECTOR GENE MC16D10L CONFERS
RESISTANCE AGAINST MELOIDOGYNE CHITWOODI IN ARABIDOPSIS AND
POTATO ...... 144
4. 1. Abstract ...... 145
4. 2. Introduction ...... 146
4. 3. Materials and methods ...... 149
4. 4. Results ...... 157
4. 5. Discussion ...... 160
vii
4. 6. Acknowledgements ...... 165
4. 7. Literature cited ...... 166
4. 8. Table ...... 177
4. 9. Figures ...... 178
CHAPTER 5: PLANT-MEDIATED RNA INTERFERENCE OF EFFECTOR GENE
MC16D10L CONFERS RESISTANCE AGAINST MELOIDOGYNE CHITWOODI IN
DIVERSE GENETIC BACKGROUNDS OF POTATO AND REDUCES
PATHOGENICITY OF NEMATODE OFFSPRING ...... 185
5. 1. Abstract ...... 186
5. 2. Introduction ...... 187
5. 3. Materials and methods ...... 191
5. 4. Results ...... 198
5. 5. Discussion ...... 203
5. 6. Acknowledgements ...... 208
5. 7. Literature cited ...... 209
5. 8. Table ...... 217
5. 9. Figures ...... 218
CHAPTER 6: BROAD MELOIDOGYNE RESISTANCE IN POTATO BASED ON
RNA INTERFERENCE OF EFFECTOR GENE 16D10 ...... 226
6. 1. Abstract ...... 227
6. 2. Introduction ...... 228
6. 3. Materials and methods ...... 230
6. 4. Results ...... 235
viii
6. 5. Discussion ...... 237
6. 6. Acknowledgments ...... 239
6. 7. Literature cited ...... 240
6. 8. Table ...... 245
6. 9. Figures ...... 246
CHAPTER 7: CONCLUSIONS ...... 253
APPENDIX ...... 258
ix
LIST OF TABLES
Chapter 3
Table S1. Primers were used in this study ...... 141
Chapter 4
Table 1. Primers and probes used for cloning, polymerase chain reaction, and northern and Southern blots ...... 177
Chapter 5
Table 1. Primers and probes used for PCR, Southern and northern blots ...... 217
Chapter 6
Table 1. Reproductive efficiency of Meloidogyne chitwoodi isolate ‘WAMC1’ on wild type and 16D10i-2 RNAi transgenic ‘Russet Burbank’ potato lines at 55 days after inoculation ...... 245
x
LIST OF FIGURES
Chapter 2
Fig. 1. Arabidopsis grown in microscopy rhizosphere chambers (micro-ROC) ...... 88
Fig. 2. Pratylenchus penetrans stained with PKH26 ...... 89
Fig. 3. Heterodera schachtii stained with PKH26 ...... 90
Fig. 4. Meloidogyne chitwoodi stained with PKH26 ...... 92
Fig. 5. Peroxisomes tagged with yellow fluorescent protein (YFP) in transgenic
Arabidopsis roots infected with Meloidogyne chitwoodi ...... 94
Fig. 6. Quantification of yellow fluorescent protein-tagged peroxisomes in
Meloidogyne chitwoodi-infected and uninfected Arabidopsis roots ...... 96
Chapter 3
Fig. 1. The localization and interaction of 16D10 in Nicotiana benthamiana ...... 134
Fig. 2. The susceptibility of Arabidopsis thaliana lines to Meloidogyne incognita ...... 135
Fig. 3. Pseudomonas syringae DC3000 infection in Arabidopsis thaliana lines ...... 136
Fig. 4. Callose deposition in Arabidopsis thaliana lines ...... 137
Fig. 5. The expression level of the PR5 gene encoding pathogen related protein 5 in
16D10-overexpressing Arabidopsis thaliana lines ...... 138
Fig. 6. The effects of overexpressing 16D10 in Arabidopsis thaliana root phenotype . 139
Fig. 7. The expression level of the VND7 gene in 16D10-overexpressing Arabidopsis thaliana lines ...... 140
Fig. S1. Yeast two-hybrid assay (Y2H) ...... 142
Fig. S2. The expression level of 16D10 gene in the 16D10-overexpressing
Arabidopsis thaliana lines ...... 143
xi
Chapter 4
Fig. 1. Sequence alignment of 16D10 orthologs from Meloidogyne incognita
(Mi16D10) and M. chitwoodi (Mc16D10L) ...... 178
Fig. 2. In situ hybridization of Mc16D10L in different Meloidogyne chitwoodi life stages ...... 178
Fig. 3. Relative transcript abundance of Mc16D10L in different Meloidogyne chitwoodi life stages ...... 179
Fig. 4. Reproductive success of Meloidogyne chitwoodi on transgenic Arabidopsis thaliana expressing pART27(16D10i-2) ...... 180
Fig. 5. Reproductive success of Meloidogyne chitwoodi on transgenic potato expressing pART27(16D10i-2) ...... 181
Fig. 6. Production of small RNAs in transgenic Arabidopsis and potato plants ...... 182
Fig. 7. Relative fold change of Mc16D10L transcript level in second-generation
Meloidogyne chitwoodi from transgenic potato lines ...... 183
Supplemental Fig. 1. Southern blot for transgenic Arabidopsis and potato ...... 184
Chapter 5
Fig. 1. Southern blots showing copy numbers of 16D10i-2 in stable transgenic potato lines ...... 218
Fig. 2. Northern blots for stable transgenic potato lines ...... 219
Fig. 3. Reproductive success of Meloidogyne chitwoodi WAMC1 on transgenic potato lines expressing 16D10i-2 ...... 220
Fig. 4. Reproductive success of Meloidogyne chitwoodi Roza on transgenic potato lines expressing 16D10i-2 in a PA99N82-4 genetic background ...... 221
xii
Fig. 5. Pathogenicity and reproductive success of M. chitwoodi offspring from potato lines with and without the 16D10i-2 RNAi transgene ...... 222
Fig. 6. Relative fold change of Mc16D10L transcript level in Meloidogyne chitwoodi second-stage juveniles (J2) offspring from different potato-nematode combinations .. 224
Chapter 6
Fig. 1. Southern blot showing copy numbers of the 16D10i-2 RNAi transgene in transformed potato lines and controls ...... 246
Fig. 2. Number of Meloidogyne chitwoodi ‘WAMC1’ eggs per plant in wild type and transgenic RNAi potato lines at 55 days after inoculation ...... 247
Fig. 3. Northern blot for 16D10i-2 RNAi transgene ...... 248
Fig. 4. Reproductive success of Meloidogyne spp. in potato lines with and without
16D10i-2 RNAi transgene ...... 249
Fig. 5. Egg masses of Meloidogyne spp. in roots of potato lines with and without
16D10i-2 RNAi transgene ...... 251
Fig. 6. Attraction and invasion of Meloidogyne incognita J2 to potato roots with and without 16D10i-2 RNAi transgene ...... 252
Appendix
Fig. 1. The effect of 16D10 in the hypersensitive responses (HR) of Nicotiana benthamiana ...... 260
xiii
CHAPTER ONE
GENERAL INTRODUCTION
1. 1. PLANT PARASITIC NEMATODES
Nematodes are non-segmented and usually vermiform worms. Phylum
Nematoda contains some of the most common and diverse animals on the Earth with more than 25,000 known species assigned to five clades. Nematodes inhabit all ecosystems and all continents (20). About 15% of described species (approximately
4,100 species) are plant parasites that cause annual yield losses of more than $80 billion (US). Thus, nematodes are one of the major threats to crop production worldwide
(44, 127). All plant parasitic nematodes range from microscopic (400 µm) to visible (8 mm) (61, 216). They have protrusible stylets (a hollow and needle-like mouth spear) to perforate cell walls, release salivary gland secretions, and take up food (88, 90).
Nematodes can be categorized by different lifestyles such as migratory, sedentary, ectoparasitic, semi-endoparasitic and endoparasitic. The top three economically important plant parasitic nematodes are root-knot nematodes
(Meloidogyne spp.), cyst nematodes (Heterodera and Globodera spp.) and root-lesion nematodes (Pratylenchus spp.) (99). All of these types are obligate parasites in host roots, totally dependent on host nutrients for survival. Even though these genera belong in the superfamily Tylenchoidea, their lifestyles are different, i.e. root-knot nematodes and cyst nematodes are sedentary while root-lesion nematodes are migratory (17).
1
1. 1. 1. Root-lesion nematodes
Root-lesion nematodes (Pratylenchus spp.) have cosmopolitan distribution and one of the widest host ranges including the following economically important crops: turfgrasses, legumes, potato, vegetables, corn, and fruit (45, 125, 176, 180-182).
Pratylenchus spp. such as P. penetrans, P. vulnus and P. coffeae reproduce bisexually while P. scribneri, P. brachyurus, P. zeae and P. neglectus are monosexual species
(147, 177). Females lay about two eggs per day in root tissues or on root surfaces
(219).
The life cycle of Pratylenchus spp. lasts from 3 to 7 weeks depending on environmental conditions and nematode species (50). The first and second-stage juveniles (J1 and J2) remain inside the egg shells. J2 hatch and molt into J3 and then
J4. Finally, J4 molt into adult males or females. All life stages except the eggs of root- lesion nematodes are infective and motile. As migratory endoparasites, during the hatched J2 to adult stages, stylets function to mechanically penetrate the elongation zone and junctures of host roots, and also facilitate migration in host roots (219). The stylets secrete cell wall-degrading enzymes that assist penetration and migration events
(186).
Each root-lesion nematode punctures and feeds on a large number of host root cells. Root-lesion nematodes use their stylets to take-up the cytoplasm of the invaded host cells, causing cell death (123). Then, these nematodes migrate to other cells to start another feeding cycle (219). Root-lesion nematodes feed from host root cells for periods ranging from minutes to hours. However, they do not form permanent feeding sites. Between the feeding and migration stages, they also have resting phases when
2 an individual nematode penetrates and coils inside one host cell for about 5 hours
(219). As a result of feeding by Pratylenchus spp, root lesions develop, and are the typical underground symptom of root-lesion nematode infection (219). Root lesions, caused by Pratylenchus spp., sometimes act as entry points for other pathogens such as Rhizoctonia fragariae, Cylindrocarpon radicicola and Verticillium dahliae. These root pathogens form disease complexes that have additional adverse effects on crop production (21, 110, 166). However, there is no specific above-ground symptom of root- lesion nematode parasitism.
1. 1. 2. Cyst nematodes
Cyst nematodes are among the most widespread and damaging nematodes in potato, wheat, soybean, sugar beet and tobacco (108). Females of these nematodes turn into cysts after they die to protect about 200 to 800 eggs inside. This is the reason these nematodes are called cyst nematodes. Cyst nematodes have several genera with both temperate and tropical occurring species. Cyst nematodes can survive without a host for long periods of time as cysts in the environment, and remain dormant for many years. Thus, it is very challenging to manage cyst nematodes by crop rotation. Cyst nematodes cause quantitative yield losses. In particular, potato cyst nematodes are responsible for about 9% of yield loss in potato production worldwide (54). Above- ground symptoms of cyst nematode infection are non-specific, and easily confused with symptoms of nutrient deficiency such as chlorosis, stunted growth and yield decline. In contrast, attached cysts on the surface of host roots are a specific underground sign of cyst nematode infection.
3
Cyst nematodes are sedentary endoparasites and obligate biotrophs. Cyst nematodes produce one to eleven generations per year depending on temperature, soil moisture, and species. The life cycle of cyst nematodes takes about 20 to 30 days to complete, starting with J1 and J2 that develop in eggshells (2, 111). Under adverse conditions or the absence of a host, J2 can remain dormant inside eggs from six to eight years (165). There are two stages of dormancy: diapause and then quiescence.
Diapause is a developmental arrest that is initiated by endogenous factors while quiescence is a developmental delay triggered by unfavorable environmental conditions
(168, 218). J2 terminate dormancy, hatch to become infective J2, and migrate toward host roots depending on host root diffusate, photoperiod, temperature and moisture (70,
121, 163).
In contrast to root lesion nematodes, only J2 are infective and only J2 have the ability to invade host roots during the cyst nematode life cycle. Infective J2 penetrate the elongation zone of host roots to become parasitic J2. Parasitic J2 migrate intra-cellularly into the vascular system of the host roots; there, they become sedentary at cells suitable for feeding sites (214). In addition to applying mechanical force, stylets release cell wall-modification enzymes that assist in penetration and migration (38, 39, 88, 89,
113, 139, 193).
At the feeding site, cyst nematodes modify both the morphology and physiology of the cortical, pericycle and protoxylem cells of the host root. Cyst nematodes transform these cells into syncytial initial cells (211, 212, 214). Hundreds of syncytial initial cells fuse together to form a syncytium, a complex feeding structure (101). The nuclei of syncytial initial cells undergo acytokinesis-endoreduplications (genomic
4 replications without cell division) and enlarge during the formation of a syncytium (40,
68). The cell walls of individual syncytial initial cells are degraded to form a cytoplasmic continuum within the syncytium (67). Syncytia provide cyst nematodes the sole source of nutrients to develop to J3 and then J4. J3 and J4 continue to feed from syncytia. J4 molt into fusiform shapes, and normally protrude from host roots. J4 molt to adult males or females. It takes about ten days to develop from J2 to adult stages, but this is temperature- and species-dependent (111). Cyst nematodes are amphimitic, so they reproduce sexually (183). Adult male cyst nematodes are non-feeding, vermiform, motile, and migrate from host roots to search and fertilize females. Females continue feeding as adults, develop saccate shapes, and produce eggs inside their bodies. As adult females fill with eggs, they die and form cysts. The cysts protect the eggs inside against adverse conditions (as reviewed by 163). Cyst size varies considerably and ranges from about 200 to 1000 micrometers (126).
There are two major genera of cyst nematodes: Heterodera and Globodera.
Heterodera spp. have brown to black, lemon-shaped cysts. The other genus,
Globodera, has spherical cysts in white, yellow or cream colors. Each cyst nematode species has a very narrow host range but causes severe diseases in important crops.
For example, H. avenae (cereal cyst nematode) only infects Gramineae and H. glycines
(soybean cyst nematode) only infects soybean and other legumes Fabaceae) (144). In contrast, H. schachtii (sugar beet cyst nematode) has a wider host range than the other species. Potato cyst nematodes (G. rostochiensis and G. pallida) and tobacco cyst nematodes (G. tabacuma) are the most important species of this genus. They mainly
5 infect Solanaceae species, including potato (Solanum tuberosum), tobacco (Nicotiana tabacum) and tomato (S. lycopersicum) (146).
1. 1. 3. Root-knot nematodes
First reported by Berkeley in 1855, root-knot nematodes (RKN, Meloidogyne spp.) are among the most destructive plant pathogens and distributed worldwide (157).
They parasitize almost all flowering plants (reviewed by 16, 184). To date, about 2,000 plant species, including a wide variety of cultivated crops, have been documented as
RKN hosts. Similar to cyst nematodes, RKN cause nonspecific above-ground symptoms such as chlorotic, stunted and distorted plants. These symptoms are normally the result of disturbed vascular systems and nutrient loss in the root systems. However, when nematodes infect underground plant organs (roots, rhizomes and tubers), they cause unique symptoms of root knots or galls.
Similar to cyst nematodes, RKN are obligate biotrophs fully depending on host plants for survival. The sedentary endoparasitic life cycle of RKN is about 21 to 35 days depending on the nematode species and the surrounding environment. The shorter the life cycle, the more efficient the parasitism. RKN proliferate fastest on suitable hosts during the growing season. The RKN life cycle starts with J1 and J2 development within eggs. J2 hatch to become infective J2, the only infective stage. J2 migrate toward the host root surface following chemical cues from root diffusates. They penetrate at the root elongation zone (the only zone where root cells increase in length) through epidermal cells and become parasitic J2 (213). Parasitic J2 migrate in an intercellular manner inside the host root toward the root tip, and then migrate back up to the root maturation zone (where root cells begin differentiation). Here, they become sedentary
6 and form feeding sites. One to two days after J2 penetration, root galls, a typical symptom of RKN infection, form, and consist of the enlarging RKN, surrounding hypertrophic giant cells, and proliferated cortical and pericycle root cells (128). Gall size varies depending on the species of RKN, the host plant, and the number of invading J2.
An exception to this is M. chitwoodi on potato where galling is undetectable to the unaided eyes.
To form feeding sites, RKN modify host vascular parenchyma root cells and turn those root cells into giant cells which then become the sole source of nutrition for the
RKN (197, 213). The giant cells are metabolically highly active and are multinucleate due to acytokinetic mitosis with repeated endoreduplication (40, 100, 204). The giant cells (normally five to eight cells/feeding site) also act as a nutrient sink for the nematode’s absorption (100). Parasitic J2 take-up nutrients from giant cells through their stylets while J3 and J4 of RKN do not feed because they are ensheathed in secondary cuticles. Adult males do not feed but regain motility when they migrate from host roots to search and fertilize adult females. Males usually only form in small numbers and under adverse environmental conditions and not in all species. In contrast, adult females remain sedentary, but they feed from giant cells to enlarge their bodies, become pyriform-shaped, and produce eggs in a gelatinous egg matrix (called egg masses) (206). Egg masses protrude from females outside the host roots and eggs are released in the environment. Most Meloidogyne spp. are parthenogenetic, meaning adult females can reproduce asexually. Parthenogenesis is an effective strategy for nematodes to strongly adapt to adverse environmental conditions by means of a high population growth rate or to overcome host resistance genes by genetic variability (29-
7
31, 184). Polyploidy of RKN genome, DNA amplification/deletion, transposable elements and different numbers of highly conserved repetitive sequences in RKN genome contribute to the genetic variability of parthenogenetic RKN (29).
Six RKN species are the most widespread and economically significant:
M. incognita, M. arenaria, M. javanica, M. hapla, M. graminicola and M. chitwoodi.
These species cause major losses for many crops such as potato, carrot, tomato, pepper, peanut and cucumber. M. hapla and M. chitwoodi are temperate RKN and are well adapted to lower temperatures while other species are distributed from tropical to sub-tropical climates (157, 175).
M. chitwoodi (Columbia RKN) was first identified in the Columbia Basin in the
Pacific Northwest (PNW) in 1980 (66). It has a relatively restricted distribution and is found throughout the western U.S., in isolated areas in Virginia, as well as in Belgium,
Argentina, Germany, the Netherlands and Turkey (as reviewed in 51). M. chitwoodi has a broad host range and infects a number of monocots and dicots, including important crops such as cereals, corn, carrots, tomato, potato and a wide range of weeds (22,
155). This feature makes crop rotation virtually ineffective as a RKN control method.
M. chitwoodi only requires a base soil temperature of 5oC to reproduce, thus making it particularly damaging in cooled vegetable storage facilities where development continues exacerbating the symptoms (138). M. chitwoodi reproduces by facultative meiotic parthenogenesis and has a high reproductive rate (138).
1. 1. 4. Molecular plant-nematode interactions
Cyst nematodes and RKN are sedentary endoparasites and among the most successful plant-parasitic nematodes because their strategy of producing secretions to
8 successfully interact with host plants is effective. These nematode secretions, normally proteins, are a vital factor in molecular plant-nematode interactions (as reviewed in 72,
77). The secretions are released from nematode openings, such as amphids, secretion- excretion pores, cuticles, phasmids and stylets. However, only effectors, protein secretions which are produced from the esophageal gland cells and secreted by nematode stylets, have been characterized thus far (38). Effectors are generally defined as proteins or small molecules produced by any pathogen that have the ability to alter the structure and function of host cells (83). Even though about 50 putative effector genes have been identified in cyst nematodes and RKN, the functions of most of the effectors have not been characterized (38). Furthermore, most of the sequences of these nematode effector genes have no similarity to genes of other organisms.
All plant-parasitic nematodes have esophageal glands. Most have three gland cells but some have up to four gland cells. Cyst nematodes and RKN have two subventral cells and one dorsal gland cell (90). In these nematodes, subventral gland cells dominate in size in infective J2 and in the early stage of parasitic J2 (90). The sizes of these cells diminish in the later stages: J3, J4 and adults. In contrast, the dorsal gland cell is smaller than the subventral cells in J2 but its size increases over time.
Thus, in the later stages, such as J4 and adults, the dorsal cell is larger than the subventral gland cells (88, 90).
Esophageal glands produce differential effectors to facilitate the different stages of nematode parasitism. Effectors whose genes are highly expressed in the subventral gland cells are thought to have crucial functions in the early stages of parasitism
(invasion, migration, and feeding site formation) (88, 90). Conversely, effector genes
9 that are highly expressed in the dorsal gland cell may play a role in the later stages of parasitism (nematode development and reproduction). These effectors are tagged by signal peptides (about 20 to 30 amino acids) at the N-terminus for exocytosis. For exocytosis, the signal peptides are cleaved and these effectors are packaged into membrane-bound granules (90). Nematode stylets, which are connected with the gland cells, inject the effector granules into the cytoplasm of host cells and some of these effectors could be transported to the apoplasm and/or nucleus of host cells during infection events (52, 87, 96, 185, 199, 217).
The functions of some putative effectors have come to light in recent years: modification of cell wall structures, suppression of host defenses, and alteration of the structure and function of host cells by targeting plant signaling pathways (77).
Nematode effector genes that encode for cell wall-modifying enzymes are well- characterized. In fact, plant-parasitic nematodes are the only animals that produce plant cell wall-modifying enzymes. The genes encoding for these enzymes are presumed to have been horizontally transferred from microbes and integrated into the nematode genome (36, 73, 98). In 1998, beta-1,4-endoglucanase was found first in cyst nematodes (H. glycines), and then in RKN (M. incognita) (150, 167, 202). After that, xylanase, pectate lyase and expansins were characterized in RKN, cyst nematodes, and root lesion nematodes (as reviewed in 73). These effectors are encoded by genes that are highly expressed in the subventral gland cells in the early stages of nematode infection. The enzymes together with the mechanical force from the stylets soften, separate, and degrade host cell walls during nematode penetration. Furthermore, RKN and cyst nematodes also can “hijack” host plant enzymes to facilitate nematode
10 parasitism (79). For example, the cellulose-binding protein (CBP), a nematode effector, binds with the plant pectin methylesterase (PME3), to elongate host roots and decrease cell wall thickness (79), thereby assisting nematode parasitism.
Besides modifying host cell walls, nematode effectors also disrupt host defense mechanisms and physiology. Chorismate mutase, a well-known nematode effector, redirects the Shikimate pathway of the host plant in order to reduce the production of salicylic acid, a key hormone in plant defense (14, 47, 85, 97, 109, 117, 120, 194). The plant salicylic acid signaling pathway also can be interrupted by an H. schachtii effector,
10A06, that binds to plant Spermidine Synthase2 (80). Hewezi et al. (80) found that overexpression of 10A06 in Arabidopsis thaliana enhanced the susceptibility of tomato not only to H. schachtii but also to Pseudomonas syringae pv. tomato (Pst DC3000). In addition, the A. thaliana plants, in which 10A06 was constitutively expressed, flowered earlier and had more leaves and longer roots than the wild type.
Rehman et al. (140) found that SPRYSEC19, another effector from
G. rostochiensis, interacted with SW5-F host protein. SW5-F gene belongs to the cluster of SW5 resistant genes in tomato (169). The interaction between SPRYSEC19 and SW5-F did not elicit the hypersensitive response (HR) in Nicotiana benthamiana, thus, this effector was thought to reduce the activity of the tomato immune system. In contrast, RBP-1, a G. pallida effector, was recognized by Gpa2, a potato resistance protein, to trigger the HR (152). Cyst nematodes also can produce the components of the ubiquitination pathway, S-phase kinase-associated protein, and ubiquitin extension proteins (77, 185). As mentioned above, these nematodes “hijack” the plant
11 ubiquitination pathway to control the protein degradation of host defenses and cell cycles.
Another function of nematode effectors is manipulating plant cell development by mimicking plant CLAVATA3 (CLV3)/ENDOSPERM SURROUNDING REGION (CLE) peptides. Cyst nematodes need the CLE-like effectors to properly form the syncytia and successfully parasitize host roots (119, 136, 141, 142, 200, 201, 203). The plant CLV3 signaling pathway regulates the differentiation and proliferation of the shoot apical meristem (105, 170). In studies by Wang et al. (201, 203), the CLE-like effectors were assumed to mimic CLV3 in order to regulate the stem cell fate of the host root. Similarly,
RKN 16D10 effector was thought to interact with SCARECROW, a plant transcription factor, to control the development of host roots (86). Of interest, RKN 16D10 has been found to be highly conserved in Meloidogyne species (86).
In response to nematode infection, host plants also have altered development and physiology through changes in gene expression. By applying micro-array, real-time
PCR, and next-generation sequencing techniques, genes encoding plant expansins, polygalacturonase, pectin acetylesterase, and cellulases have been found to be highly up-regulated at the feeding site (12, 60, 94, 174). Such enzymes loosen or degrade the host cell walls to facilitate nematode migrations and feeding site formations as reviewed by Caillaud et al. (28). The giant cells and syncytial cells also have high metabolic rates to provide water and nutrients for the nematodes. For example, the expressions of
A. thaliana transporter genes are up-regulated at the feeding sites to facilitate the transport of water and nutrients (75, 102, 112). In addition, cyst nematodes and RKN induce the production of actin and tubulin to assemble and rearrange the new
12 cytoskeleton of the feeding cell (11, 35, 41, 42). In combination with increases of cyclins and cell cycle kinases, changes in the cytoskeleton modify plant cytokinesis, cell growth, organelle movement, cell transport and cell signaling (42).
To successfully parasitize host plants, nematodes need to suppress host defense responses. This suppression has been studied through the expressions of defense marker genes. In one study, five pathogenesis-related genes (PR-1 to PR-5) were down-regulated in A. thaliana plants that were infected by M. incognita (74). These PR genes were found to be the markers of salicylic acid (SA) and jasmonic acid (JA)- dependent systemic acquired resistances (SAR) (178). When a chorismate mutase effector of M. javanica was overexpressed, both PR gene expression and salicylic acid production were reduced in A. thaliana (47). Similarly, the overexpression of 10A06, a cyst nematode effector, reduced the transcriptional levels of PR-1, PR-2 and PR-5 genes (markers of SA-dependent SAR) in A. thaliana (80). However, the expressions of
PR-3 and PR-4 genes (markers of JA-dependent SAR) were not changed. Thus, 10A06 was concluded to suppress only SA-dependent defense responses (80).
In addition, plant hormones such as ethylene, SA, and JA are thought to play defensive roles in the interaction between nematodes and host plants. SA is required for successful resistance of host plants to cyst nematodes and RKN (23, 103, 187). Defects in the production and signaling pathway of SA favor the parasitism of H. schachtii (208).
In contrast, JA has a positive role in the parasitism of nematodes. The increase of JA production or the induction of the JA signaling pathway has been shown to increase the susceptibility of tomato (15) to RKN. However, the reproduction of RKN in tomato with
JA receptors suppressed was less than in wild type plants (15). In contrast to both SA
13 and JA, the overproduction of ethylene increased the susceptibility of A. thaliana to cyst nematodes (209, 210) but reduced the susceptibility to RKN (62).
Besides ethylene, SA and JA, auxin and cytokinin are also involved in the molecular interaction between plant parasitic nematodes and host plants. Cyst nematodes have been shown to induce the accumulation of auxin in feeding sites by manipulating auxin transporters (69, 71, 112). Moreover, the auxin-responsive activities at the feeding sites of cyst nematodes were higher than those in normal root cells (1,
104). The cytokinin-responsive gene (ARR5) was induced during the initiation of RKN feeding sites but its expression was in the dividing cells surrounding giant cells (116).
1. 1. 5. Management
Plant parasitic nematodes threaten agriculture worldwide and cause losses in production of about 157 billion US dollars annually (76). In US crop production alone, more than 10 billion US dollars are lost annually due to nematode infections (65, 207).
Therefore, biological and cultural management as well as chemical control measures are all deployed to protect crop production.
Due to the significant damage that plant-parasitic nematodes cause to many important crops, current control measures rely heavily on chemical compounds, nematicides and nematistats. Nematicides directly kill nematodes while nematistats only paralyze nematodes (160). From the 19th to the first half of the 20th century, all nematicides were fumigants. Non-fumigants were first used in the second half of the
20th century (78). In the US, about 84,000 tons of chemicals, at a cost of $300 million per year, have been applied annually to manage nematode parasitism (78). To control nematodes, nematicides interfere with the key biochemical pathways in nematodes.
14
These pathways are normally highly conserved in other animals, including humans.
Therefore, nematicides are non-selective pesticides that may have adverse effects on other non-target organisms like animals and humans (78). For example, organophosphates and carbamates are especially highly toxic to birds, fish, and invertebrates. Those nematicides that are highly mobile compounds may place groundwater at risk of contamination. In addition, the nematicide methyl bromide acts as a greenhouse gas to deplete the ozone layer (78). Therefore, beginning in the 21st century, nematicides have been subjected to an increasing amount of vigorous regulations. A number of compounds, both fumigants and non-fumigants, have been phased out in recent years: chloropicrin, 1,3-dichloropropene, 1,2-dibromo-3- chloropropane, formaldehyde, carbamates, methyl bromide, organophosphates, and fenamiphos. Some chemicals, such as metam sodium, dazomet, ethoprophos, carbofuran, aldicarb, and oxamyl, still are available for controlling parasitic nematodes, but these chemicals are under review for safety reasons (92). Thus, more products are expected to be removed from the market in the near future. A few new compounds are currently being developed as nematicides and nematistats. However, it costs about 60 million US dollars over 8 to 10 years to launch a new chemical compound (78). The prediction is that fewer and fewer chemical compounds will be available to control nematode infections in crop production.
Users of synthetic nematicides are under pressure to reduce applications because of toxicity or persistence in water and the soil. Therefore biological management, using living organisms, is being considered as an alternative method to manage nematode infections (195). Taking advantage of nematode suppressive soils is
15 one tool in the biological control of nematodes. Nematode suppressive soils are commonly associated with nematophagous fungi (Pochonia chlamydosporia, Dactylella oviparasitica, and Hirsutella rhossoliensis) and bacteria (Pasteuria penetrans, Bacillus subtilis, and Pseudomonas spp.) (195). These organisms may produce compounds that inhibit the development and reproduction of nematodes. Thus, even though susceptible crops are cultivated in suppressive soils, parasitic nematode populations in these soils are significantly reduced compared to those in other soils.
Other biological control agents can be exploited to manage nematode infection
(195). Predatory nematodes, tardigrades, insects and mites are commonly used as predators of plant-parasitic nematodes. Even though these predators are abundant in natural ecosystems, the mass production and effective delivery of these predators is not feasible at the present time. Some bacterial species directly or indirectly affect the physiology and development of nematodes. Bacillus, Burkholderia and Pseudomonas species with their antibiotics, toxins, and enzymes indirectly inhibit the growth of nematode populations (134, 162). These bacteria can also enhance plant growth and defense (7, 162). In addition, Pasteuria species (P. penetrans, P. thornei, and
P. nishizawae) parasitize RKN, cyst nematodes and root-lesion nematodes, and inhibit the reproduction of these nematodes (33, 158, 171). Mass production of these bacteria has limitations as well, however.
Nematophagous trapping fungi are another common biological control agent for nematodes. Dactylella candida, Monacrosporium cionopagum, Arthrobotrys dactyloides, and Arthrobotrys oligospora produce adhesive or constricting traps to capture nematodes (10, 46, 81, 93, 159, 195). These fungi secrete toxins to immobilize
16 nematodes, and then fungal hyphae invade the nematode bodies (3, 195). Some endophytic fungi that are symbiotic with plants can improve the resistance of host plants to plant parasitic nematodes by producing nematotoxins (195). Endophytic fungi may interupt the production of root exudates that attract nematodes, but non-specifically target all nematodes, including the beneficial nematodes (predator nematodes, fungal- feeders, and bacterial-feeders). Similar to other biological control agents, nematophagous fungi are also difficult to mass produce and apply in fields. Moreover, the application of nematophagous fungi alone has low efficacy in nematode control especially in a field with a very large population size of many plant parasitic nematodes
(195). Therefore, the application of biological control agents needs to be used in combination with cultural practices.
Management of cultural practices is the time-honored technique to help prevent or eliminate plant pathogens including parasitic nematodes. Cultivation of nematode- free planting materials originating from certified nurseries is one way to prevent nematode introductions. Physical soil treatments of dry heat, steam, solar heat or flooding also can provide nematode-free growing conditions (195). Destruction of volunteer host plants, rotation of crops with varying host characteristics, allowing fields to lie fallow, using trap crops, antagonistic plants and cover crops are additional cultural practices that help prevent the build-up of parasitic nematodes (154). Further, plant parasitic nematode populations can be reduced by applying plant extracts, such as organic acids, coffee husks, phenolics, oilseed cakes, ammonia, green manure, cannabinoids, alkaloids, lactones, and mineral fertilizers (131). Sanitation during growing season and postharvest activities can help limit the spread of nematode
17 populations. Adjusting the time of planting, for example, early planting or late-sowing is also common in organic cropping systems to avoid optimal nematode infection periods
(195).
Biological control and cultural management alone are currently not sufficiently effective for long term nematode control. To date, the application of nematicides is the most effective tool to control nematode parasitism but chemical compounds are expensive and might be harmful to the environment. There is an increasing interest in finding control measures that effectively and safely control nematodes in crop production.
Breeding crops carrying natural resistance to nematodes has received much attention in agricultural research. To date, several nematode resistant (R) genes have been cloned in some crops such as sugar beet, tomato, potato, soybean, wheat, pepper, and rice. R genes that are responsible for resistance to Heterodera species have been found in sugar beet (Hs1pro-1), in rice (Hsa-1Og ) and in wheat (Cre1 and
Cre3) (43, 118, 153). To find resistant genes to Globodera species, Gpa2, Gro1-4 and
H1 were cloned from potato as well as Hero from tomato (63, 64, 132, 192).
For RKN resistance, Mi-1 is the first resistant gene that was cloned from tomato.
Plants harboring Mi-1 resist not only tropical Meloidogyne species (M. incognita,
M. javanica and M. areneria) but also whiteflies, aphids, and insects (122, 130, 148,
198). In addition, Mi-3 and Mi-9 which also were cloned from tomato, confer natural host resistance to the tropical Meloidogyne species. Moreover, Me genes that were cloned from pepper (Capsicum annuum) and Ma from plum (Prunus cerasifera) are responsible for the resistance of these crops to Meloidogyne species (18, 34).
18
Most of the resistance genes belong to the TIR-NBS-LRR, LZ-NBS-LRR and
LRR kinase groups (53, 122, 132, 205). Even though most of these R genes are ubiquitously and constitutively expressed in host plants, the expression levels of these genes are very low (179). However, the expressions of these resistance genes are induced as responses to nematode infections (179). In host plants harboring nematode resistant genes, parasitic nematodes cannot develop proper feeding sites to efficiently take-up host nutrients (143). Improperly formed feeding sites can cause an inadequate reproduction of nematode females. When nematodes penetrate resistant plants, localized cell deaths may also be observed in the host tissues surrounding nematodes
(37, 48, 137).
Cultivating nematode resistant crops is an environmentally friendly and efficient way to manage nematode infections, but there are limitations in characterizing and in deploying nematode resistant genes. One of the limitations is that A. thaliana, commonly used in cellular and molecular biology studies of flowering plants, cannot be a good model to study nematode resistance because no nematode resistant gene has been cloned from this species. Moreover, plant parasitic nematodes are unable to be directly genetically modified. Therefore, the function and mechanism of nematode resistant genes are mostly uncharacterized.
Exploiting resistant genes in nematode management is an environmentally safe approach, but it also has some disadvantages. Due to the diversity of nematode populations, the virulence of some nematode races or pathotypes may develop and overcome the resistance of R genes. For example, the RMc1(blb) gene in Solanum bulbocastanum confers resistance to M. chitwoodi race 1, but the M. chitwoodi
19 pathotype Roza breaks this gene to successfully infect S. bulbocastanum (26). In addition, it is difficult, time-consuming and labor-consuming to introduce R genes of resistant species (normally wild species) to susceptible crops by crossing species during traditional crop breeding. Moreover, crossing is sometimes impossible because of the incompatibility between different plant species. The hybrids sometimes have undesirable traits and low yields, especially when wild and domesticated species are crossed (24).
Genetic engineering is a novel approach for generating crop resistance to nematodes. Transgenic potato and A. thaliana in which snowdrop (Galanthus) lectin was expressed were found to be partially resistant to cyst nematodes and RKN (27,
145). Transgenic plants exhibiting overexpression of proteinase inhibitors also have demonstrated resistance to nematodes. Specifically, overexpression of a cysteine proteinase inhibitor (cystatin) inhibited the growth of both M. incognita and H. schachtii in A. thaliana (189). Cystatin overexpression also reduced 55% of the number of eggs of M. incognita in rice (191), as well as up to 70% of the number of eggs of M. incognita and G. pallida in potato (115). A cowpea trypsin inhibitor (CpTI) was overexpressed in
A. thaliana to inhibit the growth and reproduction of H. schachtii (190). Furthermore, the additive inhibition effect on the development and reproduction of H. schachtii was observed in transgenic A. thaliana carrying the dual overexpressed construct of cystatin and CpTI (190). In spite of these advances, protein-based genetically modified crops raise public opposition because of concerns about potential food allergies and health problems.
20
To avoid introducing a foreign protein into a plant, utilizing RNA interference
(RNAi) represents a new, alternative tactic to control parasitic nematodes. Gene silencing by the RNAi mechanism was first characterized in Caenorhabditis elegans
(59). RNAi was first applied to silence target genes of cyst nematodes in 2002 and RKN in 2005 (149, 188). The RNAi process is triggered by double-stranded RNA (dsRNA)
(59). These dsRNA are processed by the RNA-induced silencing complex (RISC) to produce short interfering RNAs (siRNA). In the RISC complex, the guide strand of siRNA binds to the target messenger RNA (mRNA) to mark this mRNA for degradation.
Thus, the translation of the target mRNA is prohibited (59). Silencing essential nematode genes with dsRNA that is complementary to the sequence of these genes can reduce nematode infections, but as important, be employed to characterize gene functions in nematode parasitism (56).
Plant parasitic nematodes can absorb dsRNA in a soaking solution through their intestines and cuticles. With current molecular technologies, though, plant parasitic nematodes cannot be stably genetically modified, which means that alternative strategies, such as transient introduction of RNAi by soaking nematodes in RNAi solutions must be used to silence the targeted nematode genes (as reviewed in 151).
The RISC pathway of nematodes can process exogenous dsRNA to trigger the silencing of target genes. To date, many studies exploit this alternative method to efficiently silence the essential genes of plant parasitic nematodes. RNAi-targeting cysteine proteinases of the intestines of G. pallida, H. glycines and M. incognita reduced the number of adult nematodes, especially females (161, 188). Other genes targeted by the RNAi soaking method are chitin synthase of nematode eggs, neuropeptides of
21 neuronal tissues, major sperm protein, C-type lectin of hypodermis, and aminopeptidase of the female reproductive system (57, 106, 114, 188). In these studies, gene silencing by RNAi soaking caused defective nematode hatches and molting as well as abnormal
J2 motility. The silencing also reduced nematode reproduction rates by increasing the ratio of males to females.
Importantly, nematode effectors, esophageal secreted proteins that play vital roles in the molecular interaction between nematodes and host plants, can be the targets of RNAi soaking. RNAi soaking mostly targets well-characterized nematode effectors, such as cellulases, pectate lyase, chorismate mutase, glutathione-S transferase, calreticulin, and polygalacturonase (8, 32, 49, 95, 149). Some RNAi targets are unknown esophageal proteins from H. glycines (8, 9); silencing in this case has resulted in the increase of the ratio of males to females and in the reduction of nematode reproduction.
Another strategy to silence nematode genes is plant-mediated RNAi (56, 215).
This approach introduces RNAi contructs into host plants. When plant parasitic nematodes feed on RNAi plants, they ingest the products of the RNAi constructs
(dsRNA or siRNA) from the host cells at feeding sites. To date, plant-mediated RNAi has successfully reduced the number of developing H. schachtii females in A. thaliana by silencing the effectors, e.g. ubiquitin-like (4G06), cellulose binding protein (3B05),
SKP1-like (8H07), zinc finger protein (10A06), and annexin-like (Hs4F01) (135, 164). In studies on the 16D10 gene, Huang et al. (84) effectively silenced this gene with the complimentary RNAi contructs, which were stably transformed into A. thaliana. Because the 16D10 encodes a highly conserved effector of Meloidogyne species (M. incognita,
22
M. javanica, M. arenaria, and M. hapla) (86), silencing of 16D10 reduced the gall numbers of these nematode species by 63 to 90%, and the egg numbers by 69 to 93%
(84). Of interest, it is not only effectors, but also other nematode genes, that are the targets of plant-mediated RNAi. RNAi constructs targeting MjTis11, integrase, Rpn7, tyrosine phosphatase, mitochondrial stress-70 protein precursor, neuropeptides of
M. incognita as well as major sperm protein of H. glycines were introduced into tobacco and soybean (56, 91, 129, 133, 172, 215). These RNAi were efficient in inhibiting the reproduction of both RKN and cyst nematodes.
In summary, plant-mediated RNAi is a promising new control strategy to develop resistant crops against a wide range of root-knot nematodes, especially by silencing effector genes. Thus, RNAi was exploited in this dissertation to generate transgenic
A. thaliana and potato plants that are tolerant of RKN.
1. 2. ROOT-KNOT NEMATODES IN POTATO
Potato, the third most economically important food crop in the world, is widely distributed worldwide in various habitats, and commonly consumed as part of a staple diet (13, 82). In 2012, potato production in the United States was 19 million metric tons compared to 374 million tons produced worldwide (5). Potato production in the Pacific
Northwest accounts for about 50% of the total production in the United States (19).
Idaho and Washington are first and second in US potato production with approximately
7 and 5 million tons, respectively, in 2012 (6).
Meloidogyne chitwoodi is a major threat to potato production in the Pacific
Northwest (55, 173). In addition, potato is also a favored host of other Meloidogyne
23 species such as M. incognita, M. hapla, M. arenaria and M. javanica. In potato,
Meloidogyne species spread through vegetative propagation of plant materials (infested seed pieces) and through cultivation practices (e.g., human, farm equipment, and irrigation). Infection by Meloidogyne species can cause serious quality defects in potato tubers (brown lesions in the flesh) and other vegetable storage organs, and can render the produce from entire fields unmarketable (58, 196). For instance, from planting to harvest, one juvenile per 250 gram soil was estimated to reproduce up to several thousand juveniles per 250 gram soil resulting in a total loss in marketability (92).
Consequently, M. chitwoodi is a strictly regulated, quarantined pest and there is no tolerance in potato export and seed potato production (4).
Soil fumigation and nonfumigant nematicides, either or both, have been used to reduce the damage caused by RKN in potato production, particularly damages due to
M. chitwoodi infection (92, 107). However, this approach is costly and environmental unfriendly. Crop rotation is not effective for controlling RKN in potato production because RKN have a wide host range (22, 155, 156). Although RMc1(blb), a resistance gene was introduced by backcrossing from a wild potato, Solanum bulbocastanum clone – SB22 (24-26, 124), there still is no commercial potato cultivar resistant to
M. chitwoodi. M. chitwoodi has two known races (WAMC1 and WAMC2) and two pathotypes (CAMC and Roza). SB22 is resistant only to WAMC1 and CAMC but susceptible to WAMC2 and Roza (25, 26). By traditional breeding, SB22 was introduced by protoplast fusion with a breeding line (R4) with the goal of introducing RMc1(blb) into cultivated potato. After a long process of multiple back crossing, an advanced breeding line PA99N82-4 was developed that has resistance against WAMC1 and CAMC, but not
24 against Roza and WAMC2 (24, 26, 124). Therefore, developing new strategies, such as interrupting the molecular interactions between nematodes and host plants by RNAi to control RKN, are crucial to maintain sustainable potato production.
1. 3. RESEARCH OBJECTIVES AND CONTRIBUTIONS
Plant-parasitic nematodes infect virtually all species of higher plants. They are a significant threat to agricultural production and global food security. Understanding plant-nematode interactions is important in the study of effective and sustainable control measures of nematode infection.
The overall goals of this dissertation were to characterize plant-nematode interaction by in vivo observation of nematode and host responses, as well as conduct functional studies of nematode effector, 16D10, and eliminate nematode infection via
RNAi. These goals were addressed by the research summarized in Chapter 2 to 6.
In Chapter 2: Nondestructive imaging of plant-parasitic nematode development and host response to nematode pathogenesis
Host root tissues normally obscure plant endoparasitic nematodes, thus, it is challenging to observe nematodes inside host roots. This study developed a novel, nondestructive technique to observe the entire process of nematode pathogenesis in planta. As proof of principle, P. penetrans, H. schachtii, and M. chitwoodi were fluorescently labeled with the lipid specific stain, PKH26, and inoculated onto A. thaliana grown in microscopy rhizosphere chambers (ROC). Nematode migration patterns and morphology were observed for the full duration of the parasitic life cycles using confocal
25 microscopy. In addition, peroxisome abundance, a host response to nematode infection, was visualized.
This study was published in Phytopathology (2014, 104:497-506) by Dinh,
P.T.Y., Knoblauch, M. and Elling, A.A. (Phytopathology Editor’s Pick in May 2014).
Phuong Dinh maintained and extracted all nematode cultures as well as generated the transgenic px-YFP A. thaliana line. She inoculated nematodes onto A. thaliana grown in
ROC, and observed living nematodes and root responses using confocal microscopy.
She also measured peroxisome abundance, performed statistical analysis and prepared the manuscript and all figures. Dr. Axel Elling and Dr. Michael Knoblauch supervised, and helped prepare the manuscript.
In Chapter 3: Meloidogyne incognita effector 16D10 modifies xylem differentiation in Arabidopsis thaliana to facilitate nematode parasitism
Nematode effectors, including 16D10, are essential for plant-nematode interactions at the molecular level, but most of their functional mechanisms have not been characterized. This study investigated the involvement of M. incognita 16D10 in the xylem development and defense system by overexpression of 16D10 in A. thaliana.
This study was prepared following the format of Molecular Plant-Microbe
Interactions. All experiments/assays in this chapter were done by Phuong Dinh. Dr. Lei
Zhang created a yeast two-hybrid (Y2H) library from cDNA of A. thaliana roots inoculated with M. incognita and a pSITE-BiFC-N1-PCK construct. Dr. Axel Elling and
Dr. Debra Inglis supervised, and helped prepare the manuscript.
26
In Chapter 4: RNA interference of effector gene Mc16D10L confers resistance against Meloidogyne chitwoodi in Arabidopsis and potato
The effector gene 16D10 is highly conserved in M. incognita, M. arenaria,
M. javanica and M. hapla, and a desirable target for RNAi to control RKN infection.
However, the existence of 16D10 in M. chitwoodi and the potential of 16D10 RNAi application for M. chitwoodi control have been unknown. Cloning 16D10 from
M. chitwoodi and studying the efficacy of 16D10 RNAi for controlling parasitism of this nematode were the objectives of this study. Stable transgenic lines of Arabidopsis and potato were generated by overexpression of dsRNA complementary to 16D10. With these transgenic plants, infection assays investigated the effect of 16D10 silencing on the reproduction of M. chitwoodi.
This study was published in Phytopathology (2014, 104:1098-1106) by Dinh,
P.T.Y., Brown, C.R. and Elling, A. A. Nematode maintenance and extraction were done by Phuong Dinh. She cloned Mc16D10L gene from cDNA of M. chitwoodi then studied the expression pattern of this gene by in situ hybridization and real-time PCR. She also generated and analyzed transgenic Arabidopsis and potato carrying 16D10 RNAi constructs by Southern and northern blots. The infection assay of M. chitwoodi on these transgenic plants and the statistical analysis also were done by Phuong Dinh. Dr. Axel
Elling and Dr. Charles Brown supervised, and helped prepare the manuscript.
27
In Chapter 5: Plant-mediated RNA interference of effector gene Mc16D10L confers resistance against Meloidogyne chitwoodi in diverse genetic backgrounds of potato and reduces pathogenicity of nematode offspring
This research investigated the effects of various potato backgrounds (cv.
Désirée, cv. Russet Burbank and PA99N82-4) on 16D10-RNAi-mediated M. chitwoodi resistance; it analyzed the efficiency of 16D10-RNAi on the reproduction of the RMc1(blb)- breaking pathotype Roza; and, studied the efficacy of 16D10 silencing on the offspring of M. chitwoodi. The 16D10-RNAi construct was transformed into selected potato backgrounds and infection assays analyzed resistance of the transgenic potato lines to
M. chitwoodi race 1 and Roza.
This study was published in Nematology (2014, 16:669-682) by Dinh, P.T.Y.,
Zhang, L., Brown, C.R. and Elling, A.A. 16D10i-2 transgenic plants of cv. Désirée and cv. Russet Burbank were generated by Phuong Dinh and transgenic PA99N82-4 potato was generated by Dr. Linhai Zhang at Prosser, WA. All Southern and northern blots were done by Phuong Dinh. She was responsible for nematode maintenance, infection assays, real-time PCR and statistical analysis in this study. Dr. Axel Elling and Dr.
Charles Brown supervised, and helped prepare the manuscript.
28
In Chapter 6: Broad Meloidogyne resistance in potato based on RNA interference of effector gene 16D10
This chapter is an extended study of the previous chapters. In this chapter, all
16D10-RNAi-transformed Russet Burbank lines were challenged for resistance against
M. chitwoodi in order to select the best line. Once identified, infection assays of
M. incognita, M. javanica, M. arenaria and M. hapla as well as nematode attraction assays were completed on potato line, D21.
This study was published in Journal of Nematology (2015, 47:71-78) by Dinh,
P.T.Y., Zhang, L., Mojtahedi, H., Brown, C.R. and Elling, A.A. Phuong Dinh generated and analyzed the 16D10i-2 Russet Burbank lines by both Southern and northern blots.
She also performed the infection assays of four Meloidogyne species (M. incognita,
M. javanica, M. arenaria, and M. hapla) on the best line, D21. She set up the nematode attraction assay and analyzed the data. The M. chitwoodi infection assay to screen the best 16D10i-2 potato line was conducted by Hassan Mojtahedi and Dr. Linhai Zhang at
Prosser, WA. Dr. Axel Elling and Dr. Charles Brown supervised and helped prepare the manuscript.
29
1. 4. Literature cited
1. Absmanner, B., Stadler, R., and Hammes, U. Z. 2013. Phloem development in
nematode-induced feeding sites: the implications of auxin and cytokinin. Front.
Plant Sci. 4:1-14.
2. Alston, D. G., and Schmitt, D. P. 1988. Development of Heterodera glycines life
stages as influenced by temperature. J. Nematol. 20:366-372.
3. Anke, H., Stadler, M., Mayer, A., and Sterner, O. 1995. Secondary metabolites
with nematicidal and antimicrobial activity from nematophagous fungi and
Ascomycetes. Can. J. Bot. 73:932-939.
4. Anonymous. 2009. Meloidogyne chitwoodi and Meloidogyne fallax. EPPO Bull.
39:5-17.
5. Anonymous. 2012. FAOSTAT. Food and Agriculture Organization of the United
Nations.
6. Anonymous. 2013. Potatoes 2012 summary. United States Department of
Agriculture/National Agricultural Statistics Service.
7. Anter, A. A., Amin, A. W., Ashoub, A. H., and El-Nuby, A. S. 2014. Evaluation of
some rhizobacteria as induce systemic resistance or bio-control agents in
controlling root-knot nematode, Meloidogyne incognita on tomato. Egypt. J.
Agronematol. 13:107-123.
8. Bakhetia, M., Urwin, P. E., and Atkinson, H. J. 2007. qPCR analysis and RNAi
define pharyngeal gland cell-expressed genes of Heterodera glycines required
for initial interactions with the host. Mol. Plant-Microbe Interact. 20:306-312.
30
9. Bakhetia, M., Urwin, P. E., and Atkinson, H. J. 2008. Characterisation by RNAi of
pioneer genes expressed in the dorsal pharyngeal gland cell of Heterodera
glycines and the effects of combinatorial RNAi. Int. J. Parasitol. 38:1589-1597.
10. Balan, J., and Lechevalier, H. A. 1972. The predaceous fungus Arthrobotrys
dactyloides: induction of trap formation. Mycologia 64:919-922.
11. Banora, M. Y., Rodiuc, N., Baldacci-Cresp, F., Smertenko, A., Bleve-Zacheo, T.,
Mellilo, M. T., Karimi, M., Hilson, P., Evrard, J. L., Favery, B., Engler, G., Abad,
P., and Engler, J. D. A. 2011. Feeding cells induced by phytoparasitic nematodes
require γ-tubulin ring complex for microtubule reorganization. PLoS Pathog.
7:e1002343.
12. Barcala, M., García, A., Cabrera, J., Casson, S., Lindsey, K., Favery, B., García-
Casado, G., Solano, R., Fenoll, C., and Escobar, C. 2010. Early transcriptomic
events in microdissected Arabidopsis nematode-induced giant cells. Plant J.
61:698-712.
13. Barrell, P. J., Meiyalaghan, S., Jacobs, J. M. E., and Conner, A. J. 2013.
Applications of biotechnology and genomics in potato improvement. Plant
Biotechnol. J. 11:907-920.
14. Bekal, S., Niblack, T. L., and Lambert, K. N. 2003. A chorismate mutase from the
soybean cyst nematode Heterodera glycines shows polymorphisms that correlate
with virulence. Mol. Plant-Microbe Interact. 16:439-446.
15. Bhattarai, K. K., Xie, Q. G., Mantelin, S., Bishnoi, U., Girke, T., Navarre, D. A.,
and Kaloshian, I. 2008. Tomato susceptibility to root-knot nematodes requires an
31
intact jasmonic acid signaling pathway. Mol. Plant-Microbe Interact. 21:1205-
1214.
16. Bird, D. M., and Kaloshian, I. 2003. Are roots special? Nematodes have their say.
Physiol. Mol. Plant Pathol. 62:115-123.
17. Blaxter, M. L., De Ley, P., Garey, J. R., Liu, L. X., Scheldeman, P., Vierstraete,
A., Vanfleteren, J. R., Mackey, L. Y., Dorris, M., Frisse, L. M., Vida, J. T., and
Thomas, W. K. 1998. A molecular evolutionary framework for the phylum
Nematoda. Nature 392:71-75.
18. Bleve-Zacheo, T., Bongiovanni, M., Melillo, M. T., and Castagnone-Sereno, P.
1998. The pepper resistance genes Me1 and Me3 induce differential penetration
rates and temporal sequences of root cell ultrastructural changes upon nematode
infection. Plant Sci. 133:79-90.
19. Bohl, W. H., and Johnson, S. B., eds. 2010. Commercial potato production in
North America. Potato Association of America Handbook. Second Revision of
American Potato Journal Supplement Volume 57 and USDA Handbook 267
20. Boucher, G., and Lambshead, P. J. D. 1995. Ecological biodiversity of marine
nematodes in samples from temperate, tropical, and deep-sea regions. Conserv.
Biol. 9:1594-1604.
21. Bowers, J. H., Nameth, S. T., Riedel, R. M., and Rowe, R. C. 1996. Infection and
colonization of potato roots by Verticillium dahliae as affected by Pratylenchus
penetrans and P. crenatus. Phytopathology 86:614-621.
22. Boydston, R. A., Mojtahedi, H., Crosslin, J. M., Brown, C. R., and Anderson, T.
2008. Effect of hairy nightshade (Solanum sarrachoides) presence on potato
32
nematodes, diseases, and insect pests. Pages 151-154 in: Annual Meeting of the
Weed-Science-Society-of-America Weed Sci Soc Amer, San Antonio, TX.
23. Branch, C., Hwang, C. F., Navarre, D. A., and Williamson, V. M. 2004. Salicylic
acid is part of the Mi-1-mediated defense response to root-knot nematode in
tomato. Mol. Plant-Microbe Interact. 17:351-356.
24. Brown, C. R., Mojtahedi, H., James, S., Novy, R. G., and Love, S. 2006.
Development and evaluation of potato breeding lines with introgressed
resistance to Columbia root-knot nematode (Meloidogyne chitwoodi). Am. J.
Potato Res. 83:1-8.
25. Brown, C. R., Mojtahedi, H., and Santo, G. S. 1989. Comparison of reproductive
efficiency of Meloidogyne chitwoodi on Solanum bulbocastanum in soil and in
vitro tests. Plant Dis. 73:957-959.
26. Brown, C. R., Mojtahedi, H., Zhang, L. H., and Riga, E. 2009. Independent
resistant reactions expressed in root and tuber of potato breeding lines with
introgressed resistance to Meloidogyne chitwoodi. Phytopathology 99:1085-
1089.
27. Burrows, P. R., and De Waele, D. 1997. Engineering resistance against plant
parasitic nematodes using anti-nematode genes. Pages 217-236 in: Cellular and
Molecular Aspects of Plant-Nematode Interactions. C. Fenoll, F. M. W. Grundler,
and S. A. Ohl, eds. Springer, Netherlands.
28. Caillaud, M. C., Dubreuil, G., Quentin, M., Perfus-Barbeoch, L., Lecomte, P., de
Almeida Engler, J., Abad, P., Rosso, M. N., and Favery, B. 2008. Root-knot
33
nematodes manipulate plant cell functions during a compatible interaction. J.
Plant Physiol. 165:104-113.
29. Castagnone-Sereno, P. 2002. Genetic variability in parthenogenetic root-knot
nematodes, Meloidogyne spp., and their ability to overcome plant resistance
genes. Nematology 4:605-608.
30. Castagnone-Sereno, P. 2006. Genetic variability and adaptive evolution in
parthenogenetic root-knot nematodes. Heredity 96:282-289.
31. Castagnone-Sereno, P., Bongiovanni, M., and Wajnberg, E. 2007. Selection and
parasite evolution: a reproductive fitness cost associated with virulence in the
parthenogenetic nematode Meloidogyne incognita. Evol. Ecol. 21:259-270.
32. Chen, Q., Rehman, S., Smant, G., and Jones, J. T. 2005. Functional analysis of
pathogenicity proteins of the potato cyst nematode Globodera rostochiensis
using RNAi. Mol. Plant-Microbe Interact. 18:621-625.
33. Chen, Z. X., and Dickson, D. W. 1998. Review of Pasteuria penetrans: Biology,
ecology, and biological control potential. J. Nematol. 30:313-340.
34. Claverie, M., Dirlewanger, E., Bosselut, N., Van Ghelder, C., Voisin, R.,
Kleinhentz, M., Lafargue, B., Abad, P., Rosso, M. N., Chalhoub, B., and
Esmenjaud, D. 2011. The Ma gene for complete-spectrum resistance to
Meloidogyne species in Prunus is a TNL with a huge repeated C-terminal post-
LRR region. Plant Physiol. 156:779-792.
35. Clément, M., Ketelaar, T., Rodiuc, N., Banora, M. Y., Smertenko, A., Engler, G.,
Abad, P., Hussey, P. J., and de Almeida Engler, J. 2009. Actin-depolymerizing
34
factor2-mediated actin dynamics are essential for root-knot nematode infection of
Arabidopsis. Plant Cell 21:2963-2979.
36. Danchin, E. G. J., Rosso, M. N., Vieira, P., de Almeida-Engler, J., Coutinho, P.
M., Henrissat, B., and Abad, P. 2010. Multiple lateral gene transfers and
duplications have promoted plant parasitism ability in nematodes. Proc. Natl.
Acad. Sci. 107:17651-17656.
37. Davies, L. J., Brown, C. R., and Elling, A. A. 2015. Calcium is involved in the
RMc1(blb)-mediated hypersensitive response against Meloidogyne chitwoodi in
potato. Plant Cell Rep. 34:167-177.
38. Davis, E. L., Hussey, R. S., and Baum, T. J. 2004. Getting to the roots of
parasitism by nematodes. Trends Parasitol. 20:134-141.
39. Davis, E. L., Hussey, R. S., Baum, T. J., Bakker, J., Schots, A., Rosso, M. N.,
and Abad, P. 2000. Nematode parasitism genes. Annu. Rev. Phytopathol.
38:365-396.
40. de Almeida Engler, J., De Vleesschauwer, V., Burssens, S., Celenza, J. L.,
Inzéc, D., Van Montagu, M., Engler, G., and Gheysen, G. 1999. Molecular
markers and cell cycle inhibitors show the importance of cell cycle progression in
nematode-induced galls and syncytia. Plant Cell 11:793-807.
41. de Almeida Engler, J., Rodiuc, N., Smertenko, A., and Abad, P. 2010. Plant actin
cytoskeleton re-modeling by plant parasitic nematodes. Plant Signal. Behav.
5:213-217.
35
42. de Almeida Engler, J., Van Poucke, K., Karimi, M., De Groodt, R., Gheysen, G.,
Engler, G., and Gheysen, G. 2004. Dynamic cytoskeleton rearrangements in
giant cells and syncytia of nematode-infected roots. Plant J. 38:12-26.
43. de Majnik, J., Ogbonnaya, F. C., Moullet, O., and Lagudah, E. S. 2003. The Cre1
and Cre3 nematode resistance genes are located at homeologous loci in the
wheat genome. Mol. Plant-Microbe Interact. 16:1129-1134.
44. Decraemer, W., and Hunt, D. J. 2006. Structure and classification. Pages 3-32 in:
Plant Nematology. R. N. Perry, and M. Moens, eds. CABI Publishing,
Wallingford, UK.
45. Dickerson, O. J., Darling, H. M., and Griffin, G. D. 1964. Pathogenicity and
population trends of Pratylenchus penetrans on potato and corn. Phytopathology
54:317-322.
46. Dowsett, J. A., and Reid, J. 1977. Transmission and scanning electron
microscope observations on the trapping of nematodes by Dactylaria candida.
Can. J. Bot. 55:2963-2970.
47. Doyle, E. A., and Lambert, K. N. 2003. Meloidogyne javanica chorismate mutase
1 alters plant cell development. Mol. Plant-Microbe Interact. 16:123-131.
48. Dropkin, V. H., Helgeson, J. P., and Upper, C. D. 1969. The hypersensitivity
reaction of tomatoes resistant to Meloidogyne incognita: reversal by cytokinins. J.
Nematol. 1:55-61.
49. Dubreuil, G., Magliano, M., Deleury, E., Abad, P., and Rosso, M. N. 2007.
Transcriptome analysis of root-knot nematode functions induced in the early
stages of parasitism. New Phytol. 176:426-436.
36
50. Duncan, L. W., and Moens, M. 2006. Migratory endoparasitic nematodes. Pages
123-152 in: Plant Nematology. R. N. Perry, and M. Moens, eds. CABI Publishing,
Wallingford, UK.
51. Elling, A. A. 2013. Major emerging problems with minor Meloidogyne species.
Phytopathology 103:1092-1102.
52. Elling, A. A., Davis, E. L., Hussey, R. S., and Baum, T. J. 2007. Active uptake of
cyst nematode parasitism proteins into the plant cell nucleus. Int. J. Parasitol.
37:1269-1279.
53. Ernst, K., Kumar, A., Kriseleit, D., Kloos, D. U., Phillips, M. S., and Ganal, M. W.
2002. The broad-spectrum potato cyst nematode resistance gene (Hero) from
tomato is the only member of a large gene family of NBS-LRR genes with an
unusual amino acid repeat in the LRR region. Plant J. 31:127-136.
54. Evans, K., and Stone, A. R. 1977. A review of the distribution and biology of the
potato cyst-nematodes Globodera rostochiensis and G. pallida. PANS 23:178-
189.
55. Evans, K., Trudgill, D. L., Raman, K. V., and Radcliffe, E. B. 1992. Pest aspects
of potato production. Part 1. The nematode pests of potatoes. Pages 438-506 in:
The Potato Crop. P. M. Harris, ed. Chapman & Hall, London, UK.
56. Fairbairn, D. J., Cavallaro, A. S., Bernard, M., Mahalinga-Iyer, J., Graham, M. W.,
and Botella, J. R. 2007. Host-delivered RNAi: an effective strategy to silence
genes in plant parasitic nematodes. Planta 226:1525-1533.
37
57. Fanelli, E., Di Vito, M., Jones, J. T., and De Giorgi, C. 2005. Analysis of chitin
synthase function in a plant parasitic nematode, Meloidogyne artiellia, using
RNAi. Gene 349:87-95.
58. Finley, A. M. 1981. Histopathology of Meloidogyne chitwoodi (Golden et al.) on
Russett Burbank potato. J. Nematol. 13:486-491.
59. Fire, A., Xu, S. Q., Montgomery, M. K., Kostas, S. A., Driver, S. E., and Mello, C.
C. 1998. Potent and specific genetic interference by double-stranded RNA in
Caenorhabditis elegans. Nature 391:806-811.
60. Fosu-Nyarko, J., Jones, M. G. K., and Wang, Z. 2009. Functional
characterization of transcripts expressed in early-stage Meloidogyne javanica-
induced giant cells isolated by laser microdissection. Mol. Plant Pathol. 10:237-
248.
61. Frederick, J. J., and Tarjan, A. C. 1989. A compendium of the genus
Pratylenchus Filipjev, 1936 (Nemata: Pratylenchidae). Revue Nématol. 12:243-
256.
62. Fudali, S. L., Wang, C., and Williamson, V. M. 2013. Ethylene signaling pathway
modulates attractiveness of host roots to the root-knot nematode Meloidogyne
hapla. Mol. Plant-Microbe Interact. 26:75-86.
63. Ganal, M. W., Simon, R., Brommonschenkel, S., Arndt, M., Phillips, M. S.,
Tanksley, S. D., and Kumar, A. 1995. Genetic mapping of a wide spectrum
nematode resistance gene (Hero) against Globodera rostochiensis in tomato.
Mol. Plant-Microbe Interact. 8:886-891.
38
64. Gebhardt, C., Mugniery, D., Ritter, E., Salamini, F., and Bonnel, E. 1993.
Identification of RFLP markers closely linked to the H1 gene conferring
resistance to Globodera rostochiensis in potato. Theor. Appl. Genet. 85:541-544.
65. Gianessi, L. P., and Carpenter, J. E. 1999. Agricultural biotechnology: Insect
control benefits. National Center for Food and Agricultural Policy, Washington
DC, USA.
66. Golden, A. M., O'Bannon, J. H., Santo, G. S., and Finley, A. M. 1980. Description
and SEM observations of Meloidogyne chitwoodi n. sp. (Meloidogynidae), a root-
knot nematode on potato in the Pacific Northwest. J. Nematol. 12:319.
67. Golinowski, W., Grundler, F. M. W., and Sobczak, M. 1996. Changes in the
structure of Arabidopsis thaliana during female development of the plant-parasitic
nematode Heterodera schachtii. Protoplasma 194:103-116.
68. Goverse, A., de Almeida Engler, J., Verhees, J., van der Krol, S., Helder, J., and
Gheysen, G. 2000. Cell cycle activation by plant parasitic nematodes. Plant Mol.
Biol. 43:747-761.
69. Goverse, A., Overmars, H., Engelbertink, J., Schots, A., Bakker, J., and Helder,
J. 2000. Both induction and morphogenesis of cyst nematode feeding cells are
mediated by auxin. Mol. Plant-Microbe Interact. 13:1121-1129.
70. Grundler, F., Schnibbe, L., and Wyss, U. 1991. In vitro studies on the behaviour
of second-stage juveniles of Heterodera schachtii (Nematoda: Heteroderidae) in
response to host plant root exudates. Parasitology 103:149-155.
39
71. Grunewald, W., Cannoot, B., Friml, J., and Gheysen, G. 2009. Parasitic
nematodes modulate PIN-mediated auxin transport to facilitate infection. PLoS
Pathog. 5:e1000266.
72. Haegeman, A., Jones, J. T., and Danchin, E. G. J. 2011. Horizontal gene transfer
in nematodes: a catalyst for plant parasitism? Mol. Plant-Microbe Interact.
24:879-887.
73. Haegeman, A., Mantelin, S., Jones, J. T., and Gheysen, G. 2012. Functional
roles of effectors of plant-parasitic nematodes. Gene 492:19-31.
74. Hamamouch, N., Li, C., Seo, P. J., Park, C. M., and Davis, E. L. 2011.
Expression of Arabidopsis pathogenesis-related genes during nematode
infection. Mol. Plant Pathol. 12:355-364.
75. Hammes, U. Z., Schachtman, D. P., Berg, R. H., Nielsen, E., Koch, W., McIntyre,
L. M., and Taylor, C. G. 2005. Nematode-induced changes of transporter gene
expression in Arabidopsis roots. Mol. Plant-Microbe Interact. 18:1247-1257.
76. Hassan, M. A., Pham, T. H., Shi, H., and Zheng, J. 2013. Nematodes threats to
global food security. Acta Agriculturae Scandinavica, Section B - Soil & Plant
Science 63:420-425.
77. Hassan, S., Behm, C. A., and Mathesius, U. 2010. Effectors of plant parasitic
nematodes that re-program root cell development. Funct. Plant Biol. 37:933-942.
78. Haydock, P. P. J., Woods, S. R., Grove, I. G., and Hare, M. C. 2006. Chemical
control of nematodes. Pages 392-410 in: Plant Nematology. R. N. Perry and M.
Moens, eds. CABI, Wallingford, UK.
40
79. Hewezi, T., Howe, P., Maier, T. R., Hussey, R. S., Mitchum, M. G., Davis, E. L.,
and Baum, T. J. 2008. Cellulose binding protein from the parasitic nematode
Heterodera schachtii interacts with Arabidopsis pectin methylesterase:
cooperative cell wall modification during parasitism. Plant Cell 20:3080-3093.
80. Hewezi, T., Howe, P. J., Maier, T. R., Hussey, R. S., Mitchum, M. G., Davis, E.
L., and Baum, T. J. 2010. Arabidopsis spermidine synthase is targeted by an
effector protein of the cyst nematode Heterodera schachtii. Plant Physiol.
152:968-984.
81. Higgins, M. L., and Pramer, D. 1967. Fungal morphogenesis: ring formation and
closure by Arthrobotrys dactyloides. Science 155:345-346.
82. Hijmans, R. J. 2001. Global distribution of the potato crop. Am. J. of Potato Res.
78:403-412.
83. Hogenhout, S. A., Van der Hoorn, R. A. L., Terauchi, R., and Kamoun, S. 2009.
Emerging concepts in effector biology of plant-associated organisms. Mol. Plant-
Microbe Interact. 22:115-122.
84. Huang, G. Z., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S. 2006.
Engineering broad root-knot resistance in transgenic plants by RNAi silencing of
a conserved and essential root-knot nematode parasitism gene. Proc. Natl. Acad.
Sci. U. S. A. 103:14302-14306.
85. Huang, G. Z., Dong, R. H., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S.
2005. Two chorismate mutase genes from the root-knot nematode Meloidogyne
incognita. Mol. Plant Pathol. 6:23-30.
41
86. Huang, G. Z., Dong, R. H., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S.
2006. A root-knot nematode secretory peptide functions as a ligand for a plant
transcription factor. Mol. Plant-Microbe Interact. 19:463-470.
87. Huang, G. Z., Gao, B. L., Maier, T., Allen, R., Davis, E. L., Baum, T. J., and
Hussey, R. S. 2003. A profile of putative parasitism genes expressed in the
esophageal gland cells of the root-knot nematode Meloidogyne incognita. Mol.
Plant-Microbe Interact. 16:376-381.
88. Hussey, R. S. 1989. Disease-inducing secretions of plant-parasitic nematodes.
Annu. Rev. Phytopathol. 27:123-141.
89. Hussey, R. S., Davis, E. L., and Baum, T. J. 2002. Secrets in secretions: genes
that control nematode parasitism of plants. Brazilian J. Plant Physiol. 14:183-194.
90. Hussey, R. S., and Mims, C. W. 1990. Ultrastructure of esophageal glands and
their secretory granules in the root-knot nematode Meloidogyne incognita.
Protoplasma 156:9-18.
91. Ibrahim, H. M. M., Alkharouf, N. W., Meyer, S. L. F., Aly, M. A. M., Gamal El-Din,
A. E. K. Y., Hussein, E. H. A., and Matthews, B. F. 2011. Post-transcriptional
gene silencing of root-knot nematode in transformed soybean roots. Exp.
Parasitol. 127:90-99.
92. Ingham, R. E., Hamm, P. B., Williams, R. E., and Swanson, W. H. 2000. Control
of Meloidogyne chitwoodi in potato with fumigant and nonfumigant nematicides.
J. Nematol. 32:556-565.
42
93. Jaffee, B. A., and Muldoon, A. E. 1995. Numerical responses of the
nematophagous fungi Hirsutella rhossiliensis, Monacrosporium cionopagum, and
M. ellipsosporum. Mycologia 87:643-650.
94. Jammes, F., Lecomte, P., Almeida-Engler, J., Bitton, F., Martin-Magniette, M. L.,
Renou, J. P., Abad, P., and Favery, B. 2005. Genome-wide expression profiling
of the host response to root-knot nematode infection in Arabidopsis. Plant J.
44:447-458.
95. Jaouannet, M., Magliano, M., Arguel, M. J., Gourgues, M., Evangelisti, E., Abad,
P., and Rosso, M. N. 2013. The root-knot nematode calreticulin Mi-CRT is a key
effector in plant defense suppression. Mol. Plant-Microbe Interact. 26:97-105.
96. Jaouannet, M., Perfus-Barbeoch, L., Deleury, E., Magliano, M., Engler, G.,
Vieira, P., Danchin, E. G. J., Rocha, M. D., Coquillard, P., Abad, P., and Rosso,
M. N. 2012. A root-knot nematode-secreted protein is injected into giant cells and
targeted to the nuclei. New Phytol. 194:924-931.
97. Jones, J. T., Furlanetto, C., Bakker, E., Banks, B., Blok, V., Chen, Q., Phillips,
M., and Prior, A. 2003. Characterization of a chorismate mutase from the potato
cyst nematode Globodera pallida. Mol. Plant Pathol. 4:43-50.
98. Jones, J. T., Furlanetto, C., and Kikuchi, T. 2005. Horizontal gene transfer from
bacteria and fungi as a driving force in the evolution of plant parasitism in
nematodes. Nematology 7:641-646.
99. Jones, J. T., Haegeman, A., Danchin, E. G. J., Gaur, H. S., Helder, J., Jones, M.
G. K., Kikuchi, T., Manzanilla-López, R., Palomares-Rius, J. E., Wesemael, W.
43
M. L., and Perry, R. N. 2013. Top 10 plant-parasitic nematodes in molecular plant
pathology. Mol. Plant Pathol. 14:946-961.
100. Jones, M. G. K. 1981. The development and function of plant cells modified by
endoparasitic nematodes. Pages 255-279 in: Plant Parasitic Nematodes. B.
Zuckerman and R. A. Rohde, eds. Academic Press, New York, USA.
101. Jones, M. G. K., and Northcote, D. H. 1972. Nematode-induced syncytium – a
multinucleate transfer cell. J. Cell Sci. 10:789-809.
102. Juergensen, K., Scholz-Starke, J., Sauer, N., Hess, P., van Bel, A. J. E., and
Grundler, F. M. W. 2003. The companion cell-specific Arabidopsis disaccharide
carrier AtSUC2 is expressed in nematode-induced syncytia. Plant Physiol.
131:61-69.
103. Kandoth, P. K., Ithal, N., Recknor, J., Maier, T., Nettleton, D., Baum, T. J., and
Mitchum, M. G. 2011. The soybean Rhg1 locus for resistance to the soybean
cyst nematode Heterodera glycines regulates expression of a large number of
stress-and defense-related genes in degenerating feeding cells. Plant Physiol.
155:1960-1975.
104. Karczmarek, A., Overmars, H., Helder, J., and Goverse, A. 2004. Feeding cell
development by cyst and root-knot nematodes involves a similar early, local and
transient activation of a specific auxin-inducible promoter element. Mol. Plant
Pathol. 5:343-346.
105. Katsir, L., Davies, K. A., Bergmann, D. C., and Laux, T. 2011. Peptide signaling
in plant development. Curr. Biol. 21:R356-R364.
44
106. Kimber, M. J., McKinney, S., McMaster, S., Day, T. A., Fleming, C. C., and
Maule, A. G. 2007. flp gene disruption in a parasitic nematode reveals motor
dysfunction and unusual neuronal sensitivity to RNA interference. FASEB J.
21:1233-1243.
107. King, B. A., and Taberna, J. J. P. 2013. Site-specific management of
Meloidogyne chitwoodi in Idaho potatoes using 1, 3-dichloropropene; approach,
experiences, and economics. J. Nematol. 45:202-213.
108. Koenning, S. R., Overstreet, C., Noling, J. W., Donald, P. A., Becker, J. O., and
Fortnum, B. A. 1999. Survey of crop losses in response to phytoparasitic
nematodes in the United States for 1994. J. Nematol. 31:587-618.
109. Lambert, K. N., Allen, K. D., and Sussex, I. M. 1999. Cloning and
characterization of an esophageal-gland-specific chorismate mutase from the
phytoparasitic nematode Meloidogyne javanica. Mol. Plant-Microbe Interact.
12:328-336.
110. LaMondia, J. A. 2003. Interaction of Pratylenchus penetrans and Rhizoctonia
fragariae in strawberry black root rot. J. Nematol. 35:17-22.
111. Lauritis, J. A., Rebois, R. V., and Graney, L. S. 1983. Development of Heterodera
glycines Ichinohe on soybean, Glycine max (L.) Merr., under gnotobiotic
conditions. J. Nematol. 15:272-281.
112. Lee, C., Chronis, D., Kenning, C., Peret, B., Hewezi, T., Davis, E. L., Baum, T. J.,
Hussey, R., Bennett, M., and Mitchum, M. G. 2011. The novel cyst nematode
effector protein 19C07 interacts with the Arabidopsis auxin influx transporter
LAX3 to control feeding site development. Plant Physiol. 155:866-880.
45
113. Lilley, C. J., Atkinson, H. J., and Urwin, P. E. 2005. Molecular aspects of cyst
nematodes. Mol. Plant Pathol. 6:577-588.
114. Lilley, C. J., Goodchild, S. A., Atkinson, H. J., and Urwin, P. E. 2005. Cloning and
characterisation of a Heterodera glycines aminopeptidase cDNA. Int. J. Parasitol.
35:1577-1585.
115. Lilley, C. J., Urwin, P. E., Johnston, K. A., and Atkinson, H. J. 2004. Preferential
expression of a plant cystatin at nematode feeding sites confers resistance to
Meloidogyne incognita and Globodera pallida. Plant Biotechnol. J. 2:3-12.
116. Lohar, D. P., Schaff, J. E., Laskey, J. G., Kieber, J. J., Bilyeu, K. D., and Bird, D.
M. 2004. Cytokinins play opposite roles in lateral root formation, and nematode
and Rhizobial symbioses. Plant J. 38:203-214.
117. Long, H., Wang, X., Xu, J. H., and Hu, Y. J. 2006. Isolation and characterization
of another cDNA encoding a chorismate mutase from the phytoparasitic
nematode Meloidogyne arenaria. Exp. Parasitol. 113:106-111.
118. Lorieux, M., Reversat, G., Diaz, S. X. G., Denance, C., Jouvenet, N., Orieux, Y.,
Bourger, N., Pando-Bahuon, A., and Ghesquière, A. 2003. Linkage mapping of
Hsa-1Og, a resistance gene of African rice to the cyst nematode, Heterodera
sacchari. Theor. Appl. Genet. 107:691-696.
119. Lu, S. W., Chen, S. Y., Wang, J. Y., Yu, H., Chronis, D., Mitchum, M. G., and
Wang, X. H. 2009. Structural and functional diversity of CLAVATA3/ESR (CLE)-
like genes from the potato cyst nematode Globodera rostochiensis. Mol. Plant-
Microbe Interact. 22:1128-1142.
46
120. Lu, S. W., Tian, D., Borchardt-Wier, H. B., and Wang, X. 2008. Alternative
splicing: a novel mechanism of regulation identified in the chorismate mutase
gene of the potato cyst nematode Globodera rostochiensis. Mol. Biochem.
Parasitol. 162:1-15.
121. Masamune, T., Anetai, M., Takasugi, M., and Katsui, N. 1982. Isolation of a
natural hatching stimulus, glycinoeclepin A, for the soybean cyst nematode.
Nature 297:495-496.
122. Milligan, S. B., Bodeau, J., Yaghoobi, J., Kaloshian, I., Zabel, P., and Williamson,
V. M. 1998. The root knot nematode resistance gene Mi from tomato is a
member of the leucine zipper, nucleotide binding, leucine-rich repeat family of
plant genes. Plant Cell 10:1307-1319.
123. Moens, M., and Perry, R. N. 2009. Migratory plant endoparasitic nematodes: A
group rich in contrasts and divergence. Annu. Rev. Phytopathol. 47:313-332.
124. Mojtahedi, H., Brown, C. R., and Santo, G. S. 1995. Characterization of
resistance in a somatic hybrid of Solanum bulbocastanum and Solanum
tuberosum to Meloidogyne chitwoodi. J. Nematol. 27:86-93.
125. Mountain, W. B., and Boyce, H. R. 1958. The peach replant problem in Ontario:
V. The relation of parasitic nematodes to regional differences in severity of peach
replant failure. Can. J. Bot. 36:125-134.
126. Mulvey, R. H. 1972. Identification of Heterodera cysts by terminal and cone top
structures. Can. J. Zool. 50:1277-1292.
127. Nicol, J. M., Turner, S. J., Coyne, D. L., den Nijs, L., Hockland, S., and Maafi, Z.
T. 2011. Current nematode threats to world agriculture. Pages 21-43 in:
47
Genomics and Molecular Genetics of Plant-Nematode Interactions. J. Jones, G.
Gheysen, and C. Fenoll, eds. Springer, Netherlands.
128. Niebel, A., De Almeida Engler, J., Hemerly, A., Ferreira, P., Inzé, D., Van
Montagu, M., and Gheysen, G. 1996. Induction of cdc2a and cyc1At expression
in Arabidopsis thaliana during early phases of nematode-induced feeding cell
formation. Plant J. 10:1037-1043.
129. Niu, J., Jian, H., Xu, J., Chen, C., Guo, Q., Liu, Q., and Guo, Y. 2012. RNAi
silencing of the Meloidogyne incognita Rpn7 gene reduces nematode parasitic
success. Eur. J. Plant Pathol. 134:131-144.
130. Nombela, G., Williamson, V. M., and Muñiz, M. 2003. The root-knot nematode
resistance gene Mi-1.2 of tomato is responsible for resistance against the
whitefly Bemisia tabaci. Mol. Plant-Microbe Interact. 16:645-649.
131. Ntalli, N. G., and Caboni, P. 2012. Botanical nematicides: a review. J. Agric.
Food Chem. 60:9929-9940.
132. Paal, J., Henselewski, H., Muth, J., Meksem, K., Menéndez, C. M., Salamini, F.,
Ballvora, A., and Gebhardt, C. 2004. Molecular cloning of the potato Gro1-4 gene
conferring resistance to pathotype Ro1 of the root cyst nematode Globodera
rostochiensis, based on a candidate gene approach. Plant J. 38:285-297.
133. Papolu, P. K., Gantasala, N. P., Kamaraju, D., Banakar, P., Sreevathsa, R., and
Rao, U. 2013. Utility of host delivered RNAi of two FMRF Amide like peptides,
flp-14 and flp-18, for the management of root knot nematode, Meloidogyne
incognita. PloS One 8:e80603.
48
134. Park, J., Seo, Y., and Kim, Y. H. 2014. Biological control of Meloidogyne hapla
using an antagonistic bacterium. Plant Pathol. J. 30:288-298.
135. Patel, N., Hamamouch, N., Li, C., Hewezi, T., Hussey, R. S., Baum, T. J.,
Mitchum, M. G., and Davis, E. L. 2010. A nematode effector protein similar to
annexins in host plants. J. Exp. Bot. 61:235-248.
136. Patel, N., Hamamouch, N., Li, C. Y., Hussey, R., Mitchum, M., Baum, T., Wang,
X. H., and Davis, E. L. 2008. Similarity and functional analyses of expressed
parasitism genes in Heterodera schachtii and Heterodera glycines. J. Nematol.
40:299-310.
137. Paulson, R. E., and Webster, J. M. 1972. Ultrastructure of the hypersensitive
reaction in roots of tomato, Lycopersicon esculentum L., to infection by the root-
knot nematode, Meloidogyne incognita. Physiol. Plant Pathol. 2:227-234.
138. Pinkerton, J. N., Santo, G. S., and Mojtahedi, H. 1991. Population dynamics of
Meloidogyne chitwoodi on Russet Burbank potatoes in relation to degree-day
accumulation. J. Nematol. 23:283-290.
139. Rehman, S., Butterbach, P., Popeijus, H., Overmars, H., Davis, E. L., Jones, J.
T., Goverse, A., Bakker, J., and Smant, G. 2009. Identification and
characterization of the most abundant cellulases in stylet secretions from
Globodera rostochiensis. Phytopathology 99:194-202.
140. Rehman, S., Postma, W., Tytgat, T., Prins, P., Qin, L., Overmars, H., Vossen, J.,
Spiridon, L. N., Petrescu, A. J., Goverse, A., Bakker, J., and Smant, G. 2009. A
secreted SPRY domain-containing protein (SPRYSEC) from the plant-parasitic
49
nematode Globodera rostochiensis interacts with a CC-NB-LRR protein from a
susceptible tomato. Mol. Plant-Microbe Interact. 22:330-340.
141. Replogle, A., Wang, J., Bleckmann, A., Hussey, R. S., Baum, T. J., Sawa, S.,
Davis, E. L., Wang, X., Simon, R., and Mitchum, M. G. 2011. Nematode CLE
signaling in Arabidopsis requires CLAVATA2 and CORYNE. Plant J. 65:430-440.
142. Replogle, A., Wang, J., Paolillo, V., Smeda, J., Kinoshita, A., Durbak, A., Tax, F.
E., Wang, X., Sawa, S., and Mitchum, M. G. 2013. Synergistic interaction of
CLAVATA1, CLAVATA2, and RECEPTOR-LIKE PROTEIN KINASE 2 in cyst
nematode parasitism of Arabidopsis. Mol. Plant-Microbe Interact. 26:87-96.
143. Rice, S. L., Stone, A. R., and Leadbeater, B. S. C. 1987. Changes in cell
structure in roots of resistant potatoes parasitized by potato cyst nematodes. 2.
Potatoes with resistance derived from Solanum vernei. Physiol. Mol. Plant
Pathol. 31:1-14.
144. Riggs, R. D. 1992. Host range. Pages 107-114 in: Biology and Management of
the Soybean Cyst Nematode. R. D. Riggs and J. A. Wrather, eds. APS Press,
Minnesota, USA.
145. Ripoll, C., Favery, B., Lecomte, P., Van Damme, E., Peumans, W., Abad, P., and
Jouanin, L. 2003. Evaluation of the ability of lectin from snowdrop (Galanthus
nivalis) to protect plants against root-knot nematodes. Plant Sci. 164:517-523.
146. Roberts, P. A., and Stone, A. R. 1981. Host ranges of Globodera species within
Solanum subgenus Leptostemonum. Nematologica 27:172-189.
147. Roman, J., and Triantaphyllou, A. C. 1969. Gametogenesis and reproduction of
seven species of Pratylenchus. J. Nematol. 1:357-362.
50
148. Rossi, M., Goggin, F. L., Milligan, S. B., Kaloshian, I., Ullman, D. E., and
Williamson, V. M. 1998. The nematode resistance gene Mi of tomato confers
resistance against the potato aphid. Proc. Natl. Acad. Sci. 95:9750-9754.
149. Rosso, M. N., Dubrana, M. P., Cimbolini, N., Jaubert, S., and Abad, P. 2005.
Application of RNA interference to root-knot nematode genes encoding
esophageal gland proteins. Mol. Plant-Microbe Interact. 18:615-620.
150. Rosso, M. N., Favery, B., Piotte, C., Arthaud, L., De Boer, J. M., Hussey, R. S.,
Bakker, J., Baum, T. J., and Abad, P. 1999. Isolation of a cDNA encoding a β -1,
4-endoglucanase in the root-knot nematode Meloidogyne incognita and
expression analysis during plant parasitism. Mol. Plant-Microbe Interact. 12:585-
591.
151. Rosso, M. N., Jones, J. T., and Abad, P. 2009. RNAi and functional genomics in
plant parasitic nematodes. Annu. Rev. Phytopathol. 47:207-232.
152. Sacco, M. A., Koropacka, K., Grenier, E., Jaubert, M. J., Blanchard, A., Goverse,
A., Smant, G., and Moffett, P. 2009. The cyst nematode SPRYSEC protein RBP-
1 elicits Gpa2-and RanGAP2-dependent plant cell death. PLoS Pathog.
5:e1000564.
153. Sandal, N. N., Salentijn, E. M. J., Kleine, M., Cai, D., Arens-de Reuver, M., Van
Druten, M., de Bock, T. S. M., Lange, W., Steen, P., Jung, C., Marcker, K.,
Stiekema, W. J., and Klein-Lankhorst, R. M. 1997. Backcrossing of nematode-
resistant sugar beet: a second nematode resistance gene at the locus containing
Hs1pro-1? Mol. Breeding 3:471-480.
51
154. Santo, G. S. 1989. The role of crop rotation systems on non-cyst nematodes
affecting potatoes in the temperate zones. Pages 121-130 in: Effects of Crop
Rotation on Potato Production in the Temperate Zones. J. Vos, C. D. Van Loon,
and G. J. Bollen, eds. Springer, Wageningen, Netherlands.
155. Santo, G. S., and O'Bannon, J. H. 1981. Pathogenicity of the Columbia root-knot
nematode (Meloidogyne chitwoodi) on wheat, corn, oat, and barley. J. Nematol.
13:548-550.
156. Santo, G. S., O'Bannon, J. H., Finley, A. M., and Golden, A. M. 1980. Occurrence
and host range of a new root-knot nematode (Meloidogyne chitwoodi) in the
Pacific Northwest. Plant Dis. 64:951-952.
157. Sasser, J. N. 1977. Worldwide dissemination and importance of the root-knot
nematodes, Meloidogyne spp. J. Nematol. 9:26-29.
158. Sayre, R. M., Wergin, W. P., Schmidt, J. M., and Starr, M. P. 1991. Pasteuria
nishizawae sp. nov., a mycelial and endospore-forming bacterium parasitic on
cyst nematodes of genera Heterodera and Globodera. Res. Microbiol. 142:551-
564.
159. Scholler, M., and Rubner, A. 1994. Predacious activity of the nematode-
destroying fungus Arthrobotrys oligospora in dependence of the medium
composition. Microbiol. Res. 149:145-149.
160. Schomaker, H. C., and Been, T. H. 2006. Plant growth and population dynamics.
Pages 275-301 in: Plant Nematology. R. N. Perry and M. Moens, eds. CABI,
Wallingford, UK.
52
161. Shingles, J., Lilley, C. J., Atkinson, H. J., and Urwin, P. E. 2007. Meloidogyne
incognita: Molecular and biochemical characterisation of a cathepsin L cysteine
proteinase and the effect on parasitism following RNAi. Exp. Parasitol. 115:114-
120.
162. Siddiqui, I. A., and Shaukat, S. S. 2002. Rhizobacteria-mediated induction of
systemic resistance (ISR) in tomato against Meloidogyne javanica. J.
Phytopathol. 150:469-473.
163. Sijmons, P. C. 1993. Plant-nematode interactions. Plant Mol. Biol. 23:917-931.
164. Sindhu, A. S., Maier, T. R., Mitchum, M. G., Hussey, R. S., Davis, E. L., and
Baum, T. J. 2009. Effective and specific in planta RNAi in cyst nematodes:
expression interference of four parasitism genes reduces parasitic success. J.
Exp. Bot. 60:315-324.
165. Slack, D. A., Riggs, R. D., and Hamblen, M. L. 1972. The effect of temperature
and moisture on the survival of Heterodera glycines in the absence of a host. J.
Nematol. 4:263-266.
166. Slootweg, A. F. G. 1956. Rootrot of bulbs caused by Pratylenchus and
Hoplolaimus spp. Nematologica 1:192-201.
167. Smant, G., Stokkermans, J. P. W. G., Yan, Y., De Boer, J. M., Baum, T. J.,
Wang, X., Hussey, R. S., Gommers, F. J., Henrissat, B., and Davis, E. L. 1998.
Endogenous cellulases in animals: isolation of β -1, 4-endoglucanase genes from
two species of plant-parasitic cyst nematodes. Proc. Natl. Acad. Sci. 95:4906-
4911.
53
168. Sommerville, R. I., and Davey, K. G. 2002. Diapause in parasitic nematodes: a
review. Can. J. Zool. 80:1817-1840.
169. Spassova, M. I., Prins, T. W., Folkertsma, R. T., Klein-Lankhorst, R. M., Hille, J.,
Goldbach, R. W., and Prins, M. 2001. The tomato gene Sw5 is a member of the
coiled coil, nucleotide binding, leucine-rich repeat class of plant resistance genes
and confers resistance to TSWV in tobacco. Mol. Breeding 7:151-161.
170. Stahl, Y., and Simon, R. 2010. Plant primary meristems: shared functions and
regulatory mechanisms. Curr. Opin. Plant Biol. 13:53-58.
171. Starr, M. P., and Sayre, R. M. 1988. Pasteuria thornei sp. nov. and Pasteuria
penetrans sensu stricto emend., mycelial and endospore-forming bacteria
parasitic, respectively, on plant-parasitic nematodes of the genera Pratylenchus
and Meloidogyne. Annales de l'Institut Pasteur/Microbiologie 139:11-31.
172. Steeves, R. M., Todd, T. C., Essig, J. S., and Trick, H. N. 2006. Transgenic
soybeans expressing siRNAs specific to a major sperm protein gene suppress
Heterodera glycines reproduction. Funct. Plant Biol. 33:991-999.
173. Strand, L. 2006. Integrated pest management for potatoes in the Western United
States. UCANR Publications.
174. Szakasits, D., Heinen, P., Wieczorek, K., Hofmann, J., Wagner, F., Kreil, D. P.,
Sykacek, P., Grundler, F. M. W., and Bohlmann, H. 2009. The transcriptome of
syncytia induced by the cyst nematode Heterodera schachtii in Arabidopsis roots.
Plant J. 57:771-784.
54
175. Taylor, A. L., Sasser, J. N., and Nelson, L. A. 1982. Relationship of climate and
soil characteristics to geographical distribution of Meloidogyne species in
agricultural soils. North Carolina State University/AID, North Carolina, USA.
176. Thies, J. A., Petersen, A. D., and Barnes, D. K. 1995. Host suitability of forage
grasses and legumes for root-lesion nematode Pratylenchus penetrans. Crop
Sci. 35:1647-1651.
177. Thistlethwayte, B. 1970. Reproduction of Pratylenchus penetrans (Nematoda:
Tylenchida). J. Nematol. 2:101-105.
178. Thomma, B. P. H. J., Eggermont, K., Penninckx, I. A. M. A., Mauch-Mani, B.,
Vogelsang, R., Cammue, B. P. A., and Broekaert, W. F. 1998. Separate
jasmonate-dependent and salicylate-dependent defense-response pathways in
Arabidopsis are essential for resistance to distinct microbial pathogens. Proc.
Natl. Acad. Sci. 95:15107-15111.
179. Thurau, T., Kifle, S., Jung, C., and Cai, D. 2003. The promoter of the nematode
resistance gene Hs1pro-1 activates a nematode-responsive and feeding site-
specific gene expression in sugar beet (Beta vulgaris L.) and Arabidopsis
thaliana. Plant Mol. Biol. 52:643-660.
180. Townshend, J. L. 1963. The pathogenicity of Pratylenchus penetrans to celery.
Can. J. Plant Sci. 43:70-74.
181. Townshend, J. L. 1963. The pathogenicity of Pratylenchus penetrans to
strawberry. Can. J. Plant Sci. 43:75-78.
182. Townshend, J. L. 1978. Infectivity of Pratylenchus penetrans on alfalfa. J.
Nematol. 10:318-323.
55
183. Triantaphyllou, A. C., and Riggs, R. D. 1979. Polyploidy in an amphimictic
population of Heterodera glycines. J. Nematol. 11:371-376.
184. Trudgill, D. L., and Blok, V. C. 2001. Apomictic, polyphagous root-knot
nematodes: exceptionally successful and damaging biotrophic root pathogens.
Annu. Rev. Phytopathol. 39:53-77.
185. Tytgat, T., Vanholme, B., De Meutter, J., Claeys, M., Couvreur, M., Vanhoutte, I.,
Gheysen, G., Van Criekinge, W., Borgonie, G., Coomans, A., and Gheysen, G.
2004. A new class of ubiquitin extension proteins secreted by the dorsal
pharyngeal gland in plant parasitic cyst nematodes. Mol. Plant-Microbe Interact.
17:846-852.
186. Uehara, T., Kushida, A., and Momota, Y. 2001. PCR-based cloning of two beta-
1,4-endoglucanases from the root-lesion nematode Pratylenchus penetrans.
Nematology 3:335-342.
187. Uehara, T., Sugiyama, S., Matsuura, H., Arie, T., and Masuta, C. 2010. Resistant
and susceptible responses in tomato to cyst nematode are differentially regulated
by salicylic acid. Plant Cell Physiol. 51:1524-1536.
188. Urwin, P. E., Lilley, C. J., and Atkinson, H. J. 2002. Ingestion of double-stranded
RNA by preparasitic juvenile cyst nematodes leads to RNA interference. Mol.
Plant-Microbe Interact. 15:747-752.
189. Urwin, P. E., Lilley, C. J., McPherson, M. J., and Atkinson, H. J. 1997.
Resistance to both cyst and root-knot nematodes conferred by transgenic
Arabidopsis expressing a modified plant cystatin. Plant J. 12:455-461.
56
190. Urwin, P. E., McPherson, M. J., and Atkinson, H. J. 1998. Enhanced transgenic
plant resistance to nematodes by dual proteinase inhibitor constructs. Planta
204:472-479.
191. Vain, P., Worland, B., Clarke, M. C., Richard, G., Beavis, M., Liu, H., Kohli, A.,
Leech, M., Snape, J., Christou, P., and Atkinson, H. 1998. Expression of an
engineered cysteine proteinase inhibitor (Oryzacystatin-IΔD86) for nematode
resistance in transgenic rice plants. Theor. Appl. Genet. 96:266-271.
192. van der Voort, J. R., Wolters, P., Folkertsma, R., Hutten, R., Van Zandvoort, P.,
Vinke, H., Kanyuka, K., Bendahmane, A., Jacobsen, E., Janssen, R., and
Bakker, J. 1997. Mapping of the cyst nematode resistance locus Gpa2 in potato
using a strategy based on comigrating AFLP markers. Theor. Appl. Genet.
95:874-880.
193. Vanholme, B., De Meutter, J., Tytgat, T., Van Montagu, M., Coomans, A., and
Gheysen, G. 2004. Secretions of plant-parasitic nematodes: a molecular update.
Gene 332:13-27.
194. Vanholme, B., Kast, P., Haegeman, A., Jacob, J., Grunewald, W. I. M., and
Gheysen, G. 2009. Structural and functional investigation of a secreted
chorismate mutase from the plant-parasitic nematode Heterodera schachtii in the
context of related enzymes from diverse origins. Mol. Plant Pathol. 10:189-200.
195. Viaene, N., Coyne, D. L., and Kerry, B. R. 2006. Biological and cultural
management. Pages 346-369 in: Plant Nematology. R. N. Perry and M. Moens,
eds. CABI, Wallingford, UK.
57
196. Viaene, N., Mahieu, T., and de la Pena, E. 2007. Distribution of Meloidogyne
chitwoodi in potato tubers and comparison of extraction methods. Nematology
9:143-150.
197. von Mende, N. 1997. Invasion and migration behaviour of sedentary nematodes.
Pages 51-64 in: Cellular and Molecular Aspects of Plant-Nematode Interactions.
C. Fenoll, F. M. W. Grundler, and S. A. Ohl, eds. Springer, Netherlands.
198. Vos, P., Simons, G., Jesse, T., Wijbrandi, J., Heinen, L., Hogers, R., Frijters, A.,
Groenendijk, J., Diergaarde, P., Reijans, M., Fierens-Onstenk, J., de Both, M.,
Peleman, J., Liharska, T., Hontelez, J., and Zabeau, M. 1998. The tomato Mi-1
gene confers resistance to both root-knot nematodes and potato aphids. Nat.
Biotechnol. 16:1365-1369.
199. Wang, J., Joshi, S., Korkin, D., and Mitchum, M. G. 2010. Variable domain I of
nematode CLEs directs post-translational targeting of CLE peptides to the
extracellular space. Plant Signal. Behav. 5:1633-1635.
200. Wang, J., Lee, C., Replogle, A., Joshi, S., Korkin, D., Hussey, R., Baum, T. J.,
Davis, E. L., Wang, X., and Mitchum, M. G. 2010. Dual roles for the variable
domain in protein trafficking and host-specific recognition of Heterodera glycines
CLE effector proteins. New Phytol. 187:1003-1017.
201. Wang, J., Replogle, A., Hussey, R., Baum, T., Wang, X., Davis, E. L., and
Mitchum, M. G. 2011. Identification of potential host plant mimics of
CLAVATA3/ESR (CLE)-like peptides from the plant-parasitic nematode
Heterodera schachtii. Mol. Plant Pathol. 12:177-186.
58
202. Wang, X., Meyers, D., Yan, Y., Baum, T., Smant, G., Hussey, R., and Davis, E.
1999. In planta localization of a β -1, 4-endoglucanase secreted by Heterodera
glycines. Mol. Plant-Microbe Interact. 12:64-67.
203. Wang, X. H., Mitchum, M. G., Gao, B. L., Li, C. Y., Diab, H., Baum, T. J., Hussey,
R. S., and Davis, E. L. 2005. A parasitism gene from a plant-parasitic nematode
with function similar to CLAVATA3/ESR (CLE) of Arabidopsis thaliana. Mol. Plant
Pathol. 6:187-191.
204. Wiggers, R. J., Starr, J. L., and Price, H. J. 1990. DNA content and variation in
chromosome number in plant cells affected by Meloidogyne incognita and M.
arenaria. Phytopathology 80:1391-1395.
205. Williamson, V. M. 1999. Plant nematode resistance genes. Curr. Opin. Plant Biol.
2:327-331.
206. Williamson, V. M., and Hussey, R. S. 1996. Nematode pathogenesis and
resistance in plants. Plant Cell 8:1735-1745.
207. Wrather, J. A., and Mitchum, M. G. 2010. Soybean cyst nematode: diagnosis and
management. University of Missouri Extension Bull.
208. Wubben, M. J. E., Jin, J., and Baum, T. J. 2008. Cyst nematode parasitism of
Arabidopsis thaliana is inhibited by salicylic acid (SA) and elicits uncoupled SA-
independent pathogenesis-related gene expression in roots. Mol. Plant-Microbe
Interact. 21:424-432.
209. Wubben, M. J. E., Rodermel, S. R., and Baum, T. J. 2004. Mutation of a UDP-
glucose-4-epimerase alters nematode susceptibility and ethylene responses in
Arabidopsis roots. Plant J. 40:712-724.
59
210. Wubben, M. J. E., Su, H., Rodermel, S. R., and Baum, T. J. 2001. Susceptibility
to the sugar beet cyst nematode is modulated by ethylene signal transduction in
Arabidopsis thaliana. Mol. Plant-Microbe Interact. 14:1206-1212.
211. Wyss, U. 1992. Observations on the feeding behaviour of Heterodera schachtii
throughout development, including events during moulting. Fund. Appl. Nematol.
15:75-89.
212. Wyss, U., and Grundler, F. M. W. 1992. Feeding-behavior of sedentary plant
parasitic nematodes. Neth. J. Plant Pathol. 98:165-173.
213. Wyss, U., Grundler, F. M. W., and Munch, A. 1992. The parasitic behaviour of
second-stage juveniles of Meloidogyne incognita in roots of Arabidopsis thaliana.
Nematologica 38:98-111.
214. Wyss, U., and Zunke, U. 1986. Observations on the behaviour of second stage
juveniles of Heterodera schachtii inside host roots. Revue Nématol. 9:153-165.
215. Yadav, B. C., Veluthambi, K., and Subramaniam, K. 2006. Host-generated
double stranded RNA induces RNAi in plant-parasitic nematodes and protects
the host from infection. Mol. Biochem. Parasitol. 148:219-222.
216. YealEs, G. W., and Van Etteger, M. 1992. Longidorus orongorongensis n. sp.
(Nematoda: Dorylaimida) from subsoil of conifer/broadleaved forest. N. Z. J.
Zool. 19:25-31.
217. Zhang, L., Davies, L. J., and Elling, A. A. 2014. A Meloidogyne incognita effector
is imported into the nucleus and exhibits transcriptional activation activity in
planta. Mol. Plant Pathol. 16:48-60.
60
218. Zheng, L., and Ferris, H. 1991. Four types of dormancy exhibited by eggs of
Heterodera schachtii. Revue Nématol. 14:419-426.
219. Zunke, U. 1990. Observations on the invasion and endoparasitic behavior of the
root lesion nematode Pratylenchus penetrans. J. Nematol. 22:309-320.
61
CHAPTER TWO
NONDESTRUCTIVE IMAGING OF PLANT-PARASITIC NEMATODE DEVELOPMENT
AND HOST RESPONSE TO NEMATODE PATHOGENESIS
Phuong T.Y. Dinh1, Michael Knoblauch2, and Axel A. Elling1
1Department of Plant Pathology, Washington State University, Pullman, WA 99164
2School of Biological Sciences, Washington State University, Pullman, WA 99164
Corresponding author : Axel A. Elling
E-mail: [email protected]
This paper was published in 2014 in Phytopathology 104 (5): 497-506
62
2. 1. ABSTRACT
The secluded lifestyle of endoparasitic plant nematodes hampers progress toward a comprehensive understanding of plant-nematode interactions. A novel technique that enables non-destructive, long-term observations of a wide range of live nematodes in planta is presented here. As proof of principle, Pratylenchus penetrans,
Heterodera schachtii and Meloidogyne chitwoodi were labeled fluorescently with PKH26 and used to infect Arabidopsis thaliana grown in microscopy rhizosphere chambers.
Nematode behavior, development, and morphology were observed for the full duration of each parasite’s life cycle by confocal microscopy for up to 27 days after inoculation.
PKH26 accumulated in intestinal lipid droplets and had no negative effect on nematode infectivity. This technique enabled visualization of Meloidogyne gall formation, nematode oogenesis, and nematode morphological features, such as the metacorpus, vulva, spicules, and cuticle. Additionally, microscopy rhizosphere chambers were used to characterize plant organelle dynamics during M. chitwoodi infection. Peroxisome abundance strongly increased in early giant cells but showed a marked decrease at later stages of feeding site development, which suggests a modulation of plant peroxisomes by root-knot nematodes during the infection process. Taken together, this technique facilitates studies aimed at deciphering plant-nematode interactions at the cellular and subcellular level and enables unprecedented insights into nematode behavior in planta.
63
2. 2. INTRODUCTION
Plant-parasitic nematodes are a major threat to sustainable crop production in temperate and tropical regions alike. Some of the most damaging species belong to the root-lesion (Pratylenchus spp.), cyst (Globodera and Heterodera spp.), and root-knot
(Meloidogyne spp.) nematodes (11,22). All plant-parasitic nematodes are obligate biotrophs that depend on their hosts for survival. However, infection strategies can show marked differences between genera. Root-lesion nematodes are migratory endoparasites that invade roots and feed on a large number of host cells that are destroyed in the process, thereby resulting in lesions on the roots (54). In contrast, sedentary endoparasites such as cyst and root-knot nematodes induce the formation of living feeding sites in infected plant tissue and minimize cellular damage to their hosts.
Syncytia and giant cells, the feeding sites of cyst and root-knot nematodes, respectively, are the sole source of nutrition for these nematodes and, therefore, essential to their survival (26). The basic principles of the life cycles of root-lesion, cyst, and root-knot nematodes are relatively similar. All four juvenile life stages (J1, J2, J3, and J4) and the development into either adult males or females are separated by molts. The first-stage juvenile (J1) remains in the egg and molts into a second-stage juvenile (J2), which hatches. In cyst and root-knot nematodes, the J2 is the infective stage that invades plant tissue and initiates the formation of a feeding site, whereas the remaining life stages (except the adult male) are sedentary (26). In root-lesion nematodes, all life stages from the J2 onward are motile and infective (54). An important difference in postinfection biology is that, in Meloidogyne spp., J3 and J4 stages do not feed because they remain ensheathed in the cuticles of the previous life stage and lose their stylets. In
64 contrast, in Heterodera spp., the cuticle of the previous life stage is completely shed and the J3 and J4 nematodes feed. In Pratylenchus spp., all vermiform life stages feed (37).
The ability to study the behavior of plant-parasitic nematodes during pathogenesis and to characterize the cellular changes in infected plant tissue is of great interest, not only to elucidate fundamental principles of host–parasite interactions but also to explore novel control strategies. Plant-parasitic nematodes secrete proteinaceous effectors from the esophageal gland cells into plant tissue to induce cellular and physiological changes in the host (33) but the basic mechanisms that define molecular and cellular plant–nematode interactions remain largely unknown. One of the reasons that significant advances in this area are slow is that endoparasitic nematodes are inherently difficult to study. As obligate biotrophs, they cannot be propagated without their hosts and, once they are deeply embedded in plant tissue, key events become obscured. One of the most commonly used methods to analyze plant– nematode relationships relies on fixing infected roots and cross-sectioning areas of interest. Depending on the research question of interest, cross-sectioning is often combined with histological or immunological staining and subsequent observations by light or fluorescence microscopy (2,38,47,48) or followed up with electron microscopy
(14,19,23,24,44). Even though this approach has yielded valuable insights, it has shortcomings that hamper conceptual breakthroughs. Cross-sectioning plant and nematode tissue is technically demanding, destructive, and prone to fixation artifacts.
Furthermore, the results give static snapshots, and do not provide a dynamic picture of cellular changes or nematode behavior over time. Whole-mounts are an alternative to cross-sections and facilitate sample preparation but are subject to the same limitations
65
(16,49). To partially overcome these problems, previous investigators have used video light microscopy (43) or fluorescent dyes to track nematode infection (15,42). However, in these cases, either the dye was lost by the nematode after only a few days or the experimental setup did not allow for in-depth in planta studies. Furthermore, most approaches are based on artificial growing conditions (i.e., plants maintained in agar or as root explants, methods that can substantially alter plant–nematode interactions).
Here, we report on an optimized technique that overcomes these challenges by fluorescently labeling root-lesion nematodes (Pratylenchus penetrans), sugar beet cyst nematodes (Heterodera schachtii), and Columbia root-knot nematodes (Meloidogyne chitwoodi) with the lipid analog PKH26 and, subsequently, following the behavior of labeled nematodes in Arabidopsis thaliana grown in microscopy rhizosphere chambers
(micro-ROC) (12) using confocal microscopy throughout each life cycle. PKH26 is a lipophilic fluorescent dye with long aliphatic tails that integrates into lipids and is widely used for long-term cell tracking assays in mammalian systems (41). PKH26 has been used to study the surface coat of nematodes (32,34,35) but its potential to observe live nematodes over longer periods in planta has not been examined to date.
2. 3. MATERIALS AND METHODS
Nematode inoculum. Nematode stock cultures were maintained under greenhouse conditions. As host plants, Mentha × piperita ‘Black Mitcham’ was used for
P. penetrans, Beta vulgaris ‘4430R’ (Betaseed, Shakopee, MN) for H. schachtii, and
Solanum lycopersicum ‘Rutgers’ for Meloidogyne chitwoodi. All host plants were grown in autoclaved sand. To obtain inoculum for experiments, mixed life stages of
66
P. penetrans were extracted by cutting infected mint roots into 1- to 2-cm pieces and shaking them in a dilute NaOCl solution (0.01% commercial bleach) on a rotary shaker at 200 rpm for 2 days at room temperature (23°C). The suspension was poured through a set of nested sieves (850-, 75-, and 25-µm pore size from top to bottom). Each sieve was consecutively rinsed with water for 2 min. Mixed life stages of P. penetrans collected on the 25-µm pore sieve were backwashed into a 50-ml tube and purified on a sucrose gradient following established procedures (17) before being used for subsequent experiments. H. schachtii cysts were collected by mixing heavily infested sand from sugar beet stock cultures with water and decanting the supernatant over a
250-µm pore sieve. Cysts were rinsed with water and crushed by manually rubbing them against the sieve surface, and the released eggs were collected on a 25-µm pore sieve. H. schachtii eggs were purified on a sucrose gradient (17) and incubated in a modified Baermann pan at room temperature (23°C) to allow hatching of J2s. The hatch solution contained 3.14 mM ZnSO4 at pH 7.0. Then, 4 to 7 days later, J2s were collected by centrifuging the hatch suspension for 3 min at 375 × g in a HN-S clinical centrifuge (International Equipment Co., Needham Heights, MA). The resulting nematode pellet was washed with sterile water and centrifuged once more before being used for PKH26 labeling. To obtain M. chitwoodi inoculum, eggs were extracted from infested tomato roots using 0.5% NaOCl (18), then purified on a sucrose gradient (17).
Eggs were rinsed with water and transferred to a modified Baermann pan to hatch J2s in sterile water at room temperature (23°C). After 4 days, J2s were collected by centrifuging the hatch suspension, as described above. Nematodes were washed with sterile water and centrifuged once more before being used for PKH26 labeling.
67
PKH26 labeling. In total, ≈50,000 mixed life stages of P. penetrans and J2s of
H. schachtii and M. chitwoodi were transferred to separate 1.5-ml microcentrifuge tubes and resuspended in 1 ml of sterile water before 1 µl of PKH26 (stock solution 1 × 10–3
M) from the MINI26 PKH26 Red Fluorescent Cell Linker Kit (Sigma-Aldrich, St. Louis,
MO) was added. Each tube was inverted several times to mix the nematode suspensions, then incubated in the dark at room temperature for 15 min. To remove excess dye, nematodes were washed five times by repeated transfers to 50 ml of sterile water followed by centrifuging at 375 × g for 3 min in a clinical centrifuge as described above. Stained nematodes were immediately used to inoculate Arabidopsis. One aliquot of each batch of PKH26-labeled inoculum was stored in water at 4°C to observe fluorescence of stained nematodes outside the host plant.
Micro-ROC. A. thaliana ecotype Col-0 seed were incubated in sterile water at
4°C for 2 days, after which seed were planted individually in micro-ROC (Advanced
Science Tools, Pullman, WA) containing moistened Sunshine number 1 potting mix
(Sun Gro Horticulture, Agawam, MA) (Fig. 1). Plants were grown in a growth room with a 14-h photoperiod (300 to 400 mEm–2 s–1) and a day-and-night temperature cycle of 20 and 15°C, respectively. Ten days after planting, when the four-leaf stage was reached, each seedling was inoculated with ≈2,000 stained nematodes by pipetting 100 µl of nematode suspension between the glass slide and the nylon mesh of each micro-ROC.
Beginning at 3 days after inoculation (DAI), plants were carefully watered and fertilized with modified Knop’s media (43) without sucrose. For each nematode species, 15 micro-ROC containing one A. thaliana seedling each were inoculated. As controls,
68 uninfected plants and unlabeled nematodes were used. All experiments were conducted twice.
Arabidopsis px-YFP line. Binary plasmid px-YFP (TAIR accession CD3-982)
(36) was introduced into Agrobacterium tumefaciens strain GV3101 and used to transform Arabidopsis thaliana ecotype Col-0 using the floral dip method (7). This created an Arabidopsis line in which all peroxisomes were labeled fluorescently by translationally fusing the peroxisomal targeting signal 1 (PTS1, Ser-Lys-Leu) to the C terminus of the yellow fluorescent protein (YFP). Transformants were selected by screening germinating seed on one-half strength Murashige & Skoog basal salt media
(Caisson Labs, North Logan, UT), supplemented with 3% D-sucrose, 0.6% Daishin agar, timentin at 0.5 g/liter, and glufosinate ammonium (Plant Media, Dublin, OH) at 10
µg/ml. Second-generation (T2) seed were used in micro-ROC experiments as described above, except that plants were inoculated with unlabeled M. chitwoodi. Fifteen micro-
ROC with one plant each were analyzed and the experiment was conducted twice.
Wild-type Arabidopsis and uninfected px-YFP plants were used as controls. For each time point (3, 8, 12, 19, and 27 DAI), the number of peroxisomes was counted in three areas of three images each in infected and uninfected roots following previously established procedures (9). Briefly, three 0.01-mm2 areas (100 by 100 µm) were cropped from each image using Adobe Photoshop. Using ImageJ, cropped RGB color images were changed to 8-bit black-and-white images. The “threshold” function was used to designate black pixels as peroxisome area. The “analyze particles” function was used to count pixel groups (number of peroxisomes) and area fraction (area percentage). This technique was unable to differentiate between overlapping
69 peroxisomes, which were counted as one organelle and possibly reduced the true number of peroxisomes in some instances. To circumvent this problem, the total area of peroxisomes was estimated ([area fraction/100] × 0.01 mm2). All data were analyzed for statistical significance using a t test at α 0.05 using SAS 9.2 software.
Confocal microscopy. Infected Arabidopsis roots and uninfected controls grown in micro-ROC were observed using an LSM 510 META laser scanning microscope
(Zeiss, Jena, Germany) beginning at 3 DAI and extending for up to 27 DAI. Excitation wavelengths were set at 514 nm (argon) and 543 nm (HeNe) with emission wavelengths of 503 to 530 nm (green) and 560 to 615 nm (red). In addition, nematodes stained with PKH26 were observed on standard microscopy glass slides 3 days after being labeled using the same parameters. For px-YFP roots and respective controls, the excitation wavelengths were set at 514 nm (argon) with the emission wavelength at
560 nm. Subsequent image processing was performed with ImageJ and Adobe
Photoshop.
2. 4. RESULTS
P. penetrans behavior and life cycle in Arabidopsis roots. Immediately after being labeled with PKH26, all vermiform life stages of P. penetrans began to show fluorescence in the lip region, esophagus, and intestine. At 3 days after labeling, pronounced staining was visible, especially in lipid droplets in the intestine of
P. penetrans that had been stored at 4°C (Fig. 2A). Increased mortality of labeled
P. penetrans or overtly toxic effects of PKH26 compared with untreated nematodes were not observed. To examine whether micro-ROC facilitates observation of root-
70 lesion nematode behavior in planta and throughout the life cycle of the parasite,
Arabidopsis grown in micro-ROC were inoculated with mixed life stages of P. penetrans that were labeled with PKH26. The nematodes migrated toward and invaded the roots as early as 1 DAI. At 3 DAI, the fluorescence of lipid droplets temporarily decreased in
PKH26-stained P. penetrans inside roots but rapidly recovered in feeding nematodes
(Fig. 2B and C). Nematodes were observed for up to 27 DAI, at which point
P. penetrans completed its life cycle. The intensity of PKH26 fluorescence was strong throughout the duration of the experiments and did not weaken at 27 DAI. Importantly, micro-ROC enabled observation of characteristic P. penetrans behavior throughout the nematode’s life cycle, including intracellular migration (Fig. 2B and C), feeding (Fig. 2C and D), coiling and resting inside host cells (Fig. 2E and F), and deposition of eggs inside host tissue (Fig. 2G and H). In addition, distinct morphological features such as the intestine, metacorpus, vulva (Fig. 2D), and spicules (Fig. 2E) were visible in labeled nematodes inside roots.
H. schachtii behavior and life cycle in Arabidopsis roots. After PKH26 and micro-ROC were used to observe the migratory endoparasite P. penetrans in planta, the technique was tested in sedentary endoparasitic nematodes. H. schachtii infective J2s were labeled and used to inoculate Arabidopsis grown in micro-ROC, as detailed above.
Similar to P. penetrans, H. schachtii infective J2s began to show PKH26-induced fluorescence immediately after being exposed to the dye. At 3 days after labeling, intestinal lipid droplets and the lip region displayed strong fluorescence in labeled J2s that were stored at 4°C (Fig. 3A). PKH26 did not have any obvious effects on the viability of J2s, which oriented themselves toward host roots, explored, and finally
71 invaded roots at the tips and in the region of cell differentiation. The fluorescence of lipid droplets temporarily decreased in nonfeeding infective J2s at 10 DAI (Fig. 3B) but quickly increased in feeding J3s at 14 DAI and even surpassed the staining intensity of
J2s in later life stages (late J3 and J4) at 22 DAI and beyond (Fig. 3C to G). In developing females, fluorescent lipid droplets decreased and almost disappeared in adult females and cysts (Fig. 3H and I). Although lipid staining decreased in adult females and cysts, residual PKH26 staining of the remaining lipid droplets, together with autofluorescence in the outer layers of the cyst, enabled a clear visualization of these important life stages. Combined with PKH26, micro-ROC enabled observation of all phases of the H. schachtii life cycle in great detail in planta, including migration through host tissue (Fig. 3B), feeding (Fig. 3C and D), molting (Fig. 3E), development of vermiform males and female cysts (Fig. 3G to I), oogenesis (Fig. 3H and I), and formation of the syncytium (Fig. 3J). Furthermore, morphological details of live
H. schachtii were clearly visible in planta, and included the esophagus, metacorpus
(Fig. 3D), intestine, (Fig. 3D to G), shed cuticle during molt (Fig. 3E), ovaries (Fig. 3H), and vulval cone (Fig. 3I).
M. chitwoodi behavior and life cycle in Arabidopsis roots. Similar to the other nematode species studied, M. chitwoodi infective J2s began to display fluorescence immediately upon exposure to PKH26. At 3 days after labeling, the lip region and lipid droplets in the intestine of J2s that were stored at 4°C showed a strong fluorescent signal (Fig. 4A). As in P. penetrans and H. schachtii, PKH26 did not have any obvious effects on the ability of M. chitwoodi infective J2s to locate and penetrate host roots. The labeled J2s migrated toward the tips of Arabidopsis roots and invaded
72 roots at the zone of cell elongation. At 3 DAI, stained J2s could be clearly observed during their intercellular migration in root tissue. The fluorescence intensity of their intestinal lipid droplets was high at 3 DAI; decreased by the time they had become parasitic J2s, when they began to induce the formation of feeding sites (Fig. 4B to F); and returned to strong intensity in late feeding life stages at 15 DAI (Fig. 4G). In contrast to developing Heterodera nematodes, which break through the root cortex, especially in roots with a small diameter such as Arabidopsis and thereby facilitate imaging,
Meloidogyne nematodes remain deeply embedded in root tissue and are surrounded by a gall. As expected, gall formation made it challenging to clearly visualize morphological details in adult M. chitwoodi females at 27 DAI, when the life cycle was complete with commencement of egg laying and observations ended (Fig. 4H and I). However, PKH26 stained the cell membranes of giant cells, which greatly facilitated nondestructive observation of feeding site morphology and development and showed the increase in giant cell size compared with infected plant cells (Fig. 4D and F).
M. chitwoodi infection modulates peroxisome abundance in giant cells in
Arabidopsis. To test whether the micro-ROC technique is useful for analyzing subcellular changes in nematode infected plant tissue, a transgenic Arabidopsis line
(px-YFP) in which the peroxisomal targeting signal 1 (PTS1, Ser-Lys-Leu) is fused to the C terminus of YFP and, thereby, fluorescently labeling peroxisomes (36), was inoculated with M. chitwoodi infective J2s. Root tissue peroxisomes were evenly distributed throughout uninfected roots (Fig. 5). At 3 DAI, M. chitwoodi initiated feeding site formation and the number and area of peroxisomes per cell significantly increased in infected root cells (Figs. 5 and 6). At 8 DAI, the number of peroxisomes was higher in
73 infected plant cells when observed with the microscope (Fig. 5). However, due to technical constraints that did not allow differentiating single peroxisomes that overlapped in the microscope image, the counting technique used indicated that there was no statistically significant difference in the number of peroxisomes between infected and uninfected cells at 8 DAI (Fig. 6A). Measuring the area of peroxisomes was not affected by these technical constraints (see Materials and Methods) and showed a significant increase in the area of peroxisomes in infected cells at 8 DAI (Fig. 6B). At later stages of the infection cycle, the relative distribution of peroxisomes reversed. At
12 DAI, there were significantly fewer peroxisomes in giant cells than in the surrounding plant cells and uninfected areas. This trend continued throughout the remaining time points at 19 and 27 DAI, when the life cycle of M. chitwoodi was complete and observations ended (Fig. 5D to G). Taken together, we found that the number of peroxisomes increased in young giant cells compared with uninfected plant tissue whereas, in later stages of the M. chitwoodi life cycle, there were significantly fewer peroxisomes in nematode feeding sites relative to in neighboring uninfected plant tissue.
2. 5. DISCUSSION
Attempts to study live nematodes in their hosts go back to the early 1940s with the pioneering experiments of Linford, who developed root observation boxes to observe Meloidogyne spp. (27,28). Even though this method allowed root development in soil, thereby having the advantage of offering relatively natural growing conditions,
Linford’s design did not enable prolonged in planta studies. Early in planta nematode
74 behavior could be imaged inside the root observation boxes but nematode migration events and later behavior could only be followed in dissected roots, which limited the usefulness of this technique (28). Significant breakthroughs in plant–nematode imaging were made by growing Arabidopsis in agar under axenic conditions, infecting plants with surface-sterilized nematodes, and observing the resulting interactions with video- enhanced contrast light microscopy (14,43,50,51). A major advantage of this approach was that it was nondestructive and enabled long-term observations of live nematodes in planta. Subsequent studies made use of fluorescent dyes or plant reporter genes to visualize details of live plant–nematode interactions (6,45). However, even minor changes in the composition of artificial plant growth media can have a major impact on the infection rate of nematodes and, therefore, alter the relationships with the host, which creates a major challenge when using axenic growth conditions (43).
Furthermore, imaging of live nematodes during the infection process would greatly benefit from a fluorescent label that could be applied to the parasite and facilitate following its behavior in planta. Plant-parasitic nematodes are known to take up fluorescent dyes such as acridine orange, fluorescein isothiocyanate, or fluorescein diacetate (4,5,15,42). However, these dyes proved to be of relatively little value in plant–nematode interaction studies because they were not stable throughout the entire life of the nematode, thereby obscuring late parasitic events (15,42).
To resolve these problems, an optimized method to observe plant–nematode interactions based on fluorescently labeling nematodes with PKH26 and following behavior inside roots using plants grown in micro-ROC was developed. The technique has distinct advantages over previously used imaging methods. Most importantly, it
75 enables nondestructive long-term observations of plant–nematode interactions over the entire life of the parasite. Furthermore, host plants are grown in potting soil and roots are not exposed to light. These growing conditions sidestep potential problems with media composition that can influence the ability of nematodes to infect (43) and avoid the need to surface-sterilize nematodes with compounds such as mercuric chloride, which is lethal at high concentrations and can have a negative impact on host–parasite interactions, even at low concentrations, by interfering with nematode olfaction, neurotransmission, and locomotion (31). Roots grown with exposure to light show a greater total length, more branching, more root hairs, and bursts of reactive oxygen species (52,53), traits that can influence nematode infectivity. Additionally, exposure of nematode-infected root systems to light is known to cause the formation of chloroplasts in feeding sites of sedentary endoparasites (40,45). Recent studies provide evidence for molecular cross-talk between photosynthesis, plant immunity, and chloroplast-derived reactive oxygen species involved in defense responses (13,20,55). Plant–nematode interaction studies based on culture conditions that permit permanent light exposure of roots grown in agar are altering the relationships between the nematode and its host, thereby possibly inviting systematic errors and confounding variables that can easily be avoided with the technique presented here. In this study, we have limited our observations to A. thaliana due to its ease of culture but micro-ROC have been used to study root biology in a wide variety of plants, including tobacco, wheat, and onion (M.
Knoblauch, unpublished data). The versatility of the micro-ROC growing system and the ability of PKH26 to stain a wide range of plant-parasitic nematodes suggest that this technique can be applied in diverse plant–nematode pathosystems.
76
PKH26 labeling was extremely stable for the duration of the life cycle of all three nematode species tested here. The dye accumulated predominantly in the intestinal lipid granules and, to a lesser degree, in the surface coat of nematodes. In our experiments, we saw no obvious change in the behavior and biology of the nematodes stained with PKH26 and, thus, have no reason to believe that other aspects of the nematodes’ biology are affected by the staining. In nonfeeding J2s of H. schachtii and
M. chitwoodi, the intensity of PKH26 fluorescence temporarily decreased but, once feeding commenced and the intestinal lipid reservoirs were replenished, fluorescence returned to high intensity in juveniles and young adults before it decreased again in adults. This pattern is consistent with the changes in lipid concentration in developing nematodes and the abundance of lipid droplets in the intestine (1,10,25,30).
Furthermore, lipid droplets are dynamic and can separate into smaller droplets or merge into larger ones (39), which suggests that the observed changes in PKH26 fluorescence throughout feeding and nonfeeding nematode life stages mirror this dynamic by incorporating stain from older, previously labeled lipid droplets into newly formed ones.
Nematodes were labeled fluorescently almost immediately upon PKH26 exposure, which points at a diffusion mechanism across cell membranes rather than ingestion of the dye. However, PKH26 was unable to cross the nematode egg shell and did not label developing juveniles inside eggs (data not shown). Interestingly, we found that the giant cells of PKH26-labeled root-knot nematodes also displayed PKH26-specific fluorescence. Giant cells are known to develop massive ingrowths of the plasma membrane, which increase their internal surface area 10 times or more and lead to a corresponding increase in membrane lipids (23). Thus, it is feasible that the giant-cell-
77 specific fluorescence observed here results from incorporation of PKH26 that was released into the feeding site by the nematode. In control plants that were infected with nonlabeled nematodes, no comparable fluorescence could be observed.
To validate the use of micro-ROC to image nematode-induced changes in plant cells during the infection process, the response of peroxisomes to root-knot nematode infection was studied. We found that the number of plant peroxisomes strongly increased in early giant cells as early as 3 DAI compared with uninfected cells. At later stages of the infection process, peroxisomes were significantly less abundant in giant cells than in uninfected plant cells. Peroxisomes are found in all eukaryotes and are involved in both reactive oxygen species decomposition and production (8). Hydrogen peroxide, a universal cell stress signal, can induce the formation of new peroxisomes in plants and animals (29). The changes in peroxisome abundance in root-knot nematode- infected plant cells observed in this study mirror changes in the transcript level of peroxisome-specific genes in infected plants. Using differential display assays,
Vercauteren and co-authors (46) showed that the Arabidopsis peroxidase gene ATP6a was induced in young giant cells and endodermal cells at 3 to 7 DAI but was restricted to endodermal cells at later time points between 14 and 21 DAI, which corresponds to the location of peroxisomes observed here. Similarly, Arabidopsis peroxidase and catalase genes were found to be downregulated in giant cells in large-scale microarray experiments (3,21). We hypothesize that root-knot nematodes induce, directly or indirectly, a degradation of peroxisomes in developing giant cells to protect their feeding sites from peroxisome-derived reactive oxygen species involved in plant defense responses.
78
In summary, the combined use of micro-ROC and fluorescent labels has the potential to significantly advance fundamental understanding of plant–nematode interactions. Further, it has broad appeal for studying rhizosphere biology in general when applied to other plant-associated organisms such as bacteria, oomycetes, or fungi.
2. 6. ACKNOWLEDGMENTS
This research was supported by funding from the Washington State Department of Agriculture, Washington State Potato Commission, Idaho Potato Commission, and
Washington Grain Commission to A.A.E. PPNS Number 0622, Department of Plant
Pathology, College of Agricultural, Human, and Natural Resource Sciences, Agricultural
Research Center, Project Number WNP00744, Washington State University, Pullman
99164- 6430. We thank C. Davitt and V. Lynch-Holm (Franceschi Microscopy & Imaging
Center, Washington State University) for excellent technical assistance with confocal microscopy.
79
2. 7. LITERATURE CITED
1. Atkinson, H. J., and Harris, P. D. 1988. Changes in nematode antigens
recognized by monoclonal antibodies during early infections of soya beans with
the cyst nematode Heterodera glycines. Parasitology 98:479-487.
2. Banora, M. Y., Rodiuc, N., Baldacci-Cresp, F., Smertenko, A., Bleve-Zacheo, T.,
Mellilo, M. T., Karimi, M., Hilson, P., Evrand, J. L., Favery, B., Engler, G., Abad,
P., and de Almeida-Engler, J. 2011. Feeding cells induced by phytoparasitic
nematodes require γ-tubulin ring complex for microtubule reorganization. PLoS
Pathog. 7:e1002343.
3. Barcala, M., García, A., Cabrera, J., Casson, S., Lindsey, K., Favery, B., García-
Casado, G., Solano, R., Fenoll, C., and Escobar, C. 2010. Early transcriptomic
events in microdissected Arabidopsis nematode-induced giant cells. Plant J.
61:698-712.
4. Bird, A. F. 1979. A method of distinguishing between living and dead nematodes
by enzymatically induced fluorescence. J. Nematol. 11:103-105.
5. Blair, L., Perry, R. N., Oparka, K., and Jones, J. T. 1999. Activation of
transcription during the hatching process of the potato cyst nematode Globodera
rostochiensis. Nematology 1:103-111.
6. Böckenhoff, A., Prior, D. A. M., Grundler, F. M. W., and Oparka, K. J. 1996.
Induction of phloem unloading in Arabidopsis thaliana roots by the parasitic
nematode Heterodera schachtii. Plant Physiol. 112:1421-1427.
80
7. Clough, S. J., and Bent, A. F. 1998. Floral dip: a simplified method for
Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16:735-
743.
8. del Río, L. A., Corpas, F. J., Sandalio, L. M., Palma, J. M., Gómez, M., and
Barroso, J. B. 2002. Reactive oxygen species, antioxidant systems and nitric
oxide in peroxisomes. J. Exp. Bot. 53:1255-1272.
9. Desai, M., and Hu, J. 2008. Light induces peroxisome proliferation in Arabidopsis
seedlings through the photoreceptor phytochrome A, the transcription factor HY5
HOMOLOG, and the peroxisomal protein PEROXIN11b. Plant Physiol. 146:1117-
1127.
10. Dropkin, V. H., and Acedo, J. 1974. An electron microscopic study of glycogen
and lipid in female Meloidogyne incognita (root-knot nematode). J. Parasitol.
60:1013-1021.
11. Elling, A. A. 2013. Major emerging problems with minor Meloidogyne species.
Phytopathology 103:1092-1102.
12. Froehlich, D. R., Mullendore, D. L., Jensen, K. H., Ross-Elliott, T. J., Anstead, J.
A., Thompson, G. A., Pélissier, H. C., and Knoblauch, M. 2011. Phloem
ultrastructure and pressure flow: sieve-element-occlusion-related agglomerations
do not affect translocation. Plant Cell 23:4428-4445.
13. Göhre, V., Jones, A. M. E., Sklenár, J., Robatzek, S., and Weber, A. P. M. 2012.
Molecular crosstalk between PAMP-triggered immunity and photosynthesis. Mol.
Plant-Microbe Interact. 25:1083-1092.
81
14. Golinowski, W., Grundler, F. M. W., and Sobczak, M. 1996. Changes in the
structure of Arabidopsis thaliana during female development of the plant-parasitic
nematode Heterodera schachtii. Protoplasma 194:103-116.
15. Goto, D. B., Fosu-Nyarko, J., Sakuma F., Sadler, J, Flottman-Reid, M., Uehara,
T., Kondo, N., Yamaguchi, J., and Jones, M. G. K. 2010. In planta observation of
live fluorescent plant endoparasitic nematodes during early stages of infection.
Nematol. Res. 40:15-19.
16. Guhl, K. E., Czymmek, K., and Sherrier, D. J. 2004. Multi-photon microscopy as
a tool to track nematode infection. Microsc. Microanal. 10 (Suppl.):214-215.
17. Humphreys-Pereira, D. A., and Elling, A. A. 2014. Morphological analysis of
males and second-stage juveniles of Meloidogyne chitwoodi. Nematology
16:149-162.
18. Hussey, R. S., and Barker, K. R. 1973. A comparison of methods of collecting
inocula of Meloidogyne spp., including a new technique. Plant Dis. Rep. 57:1025-
1028.
19. Hussey, R. S., and Mims, C. W. 1991. Ultrastructure of feeding tubes formed in
giant-cells induced in plants by the root-knot nematode Meloidogyne incognita.
Protoplasma 162:99-107.
20. Islam, S. Z., Baradoost, M., Bekal, S., and Lambert, K. 2008. Red light-induced
systemic disease resistance against root-knot nematode Meloidogyne javanica
and Pseudomonas syringae pv. tomato DC 3000. J. Phytopathol. 156:708-714.
21. Jammes, F., Lecomte, P., de Almeida-Engler, J., Bitton, F., Martin-Magniette, M.
L., Renou, J. P., Abad, P., and Favery, B. 2005. Genome-wide expression
82
profiling of the host response to root-knot nematode infection in Arabidopsis.
Plant J. 44:447-458.
22. Jones, J. T., Haegeman, A., Danchin, E. G. J., Gaur, H. S., Helder, J., Jones, M.
G. K., Kikuchi, T., Manzanilla-López, R., Palomares-Rius, J. E., Wesemael, W.
M. L., and Perry, R. N. 2013. Top 10 plant-parasitic nematodes in molecular plant
pathology. Mol. Plant Pathol. 14:946-961.
23. Jones, M. G. K., and Dropkin, V. H. 1976. Scanning electron microscopy of
nematode-induced giant transfer cells. Cytobios 15:149-161.
24. Jones, M. G. K., and Northcote, D. H. 1972. Multinucleate transfer cells induced
in Coleus roots by the root-knot nematode, Meloidogyne arenaria. Protoplasma
75:381-395.
25. Krusberg, L. R., Hussey, R. S., and Fletcher, C. L. 1973. Lipid and fatty acid
composition of females and eggs of Meloidogyne incognita and M. arenaria.
Comp. Biochem. Physiol. 45:335-341.
26. Kyndt, T., Vieira, P., Gheysen, G., and de Almeida-Engler, J. 2013. Nematode
feeding sites: unique organs in plant roots. Planta 238:807-818.
27. Linford, M. B. 1940. A miniature root-observation box. Phytopathology 30:348-
349.
28. Linford, M. B. 1942. The transient feeding of root-knot nematode larvae.
Phytopathology 32:580-589.
29. Lopez-Huertas, E., Charlton, W. L., Johnson, B., Graham, I. A., and Baker, A.
2000. Stress induces peroxisome biogenesis genes. EMBO J. 19:6770-6777.
83
30. Marwah, R., and Khera, S. 1985. Histochemical localization of proteins,
carbohydrates and lipids in females of Meloidogyne incognita. Riv. Parasitol.
46:313-322.
31. McElwee, M. K., and Freedman, J. H. 2011. Comparative toxicology of
mercurials in Caenorhabditis elegans. Environ. Toxicol. Chem. 30:2135-2141.
32. Mendoza, M. E. L. D., Modha, J., Roberts, M. C., Curtis, R., and Kusel, J. R.
2000. Changes in the lipophilicity of the surfaces of Meloidogyne incognita and
Haemonchus contortus during exposure to host signals. Parasitology 120:203-
209.
33. Mitchum, M. G., Hussey, R. S., Baum, T. J., Wang, X., Elling, A. A., Wubben, M.,
and Davis, E. L. 2013. Nematode effector proteins: an emerging paradigm of
parasitism. New Phytol. 199:879-894.
34. Modha, J., Kusel, J. R., and Kennedy, M. W. 1995. A role for second
messengers in the control of activation-associated modification of the surface of
Trichinella spiralis infective larvae. Mol. Biochem. Parasitol. 72:141-148.
35. Modha, J., Piedrafita, D., Roberts, M. C., Kusel, J. R., and Kennedy, M. W. 2000.
Secretion of IL-12 by murine macrophages activated by immunoglobulin
receptor-mediated internalization of the surface coat of Trichinella spiralis larvae.
Parasite Immunol. 22:115-120.
36. Nelson, B. K., Cai, X., and Nebenführ, A. 2007. A multicolored set of in vivo
organelle markers for co-localization studies in Arabidopsis and other plants.
Plant J. 51:1126-1136.
84
37. Nickle, W. R., ed. 1991. Manual of Agricultural Nematology. Marcel Dekker Inc.,
New York.
38. Nobre, M. J. G., von Mende, N., Dolan, L., Schmidt, K. P., Evans, K., and
Mulligan, B. 1995. Immunolabeling of cell surfaces of Arabidopsis thaliana roots
following infection by Meloidogyne incognita. J. Exp. Bot. 46:1711-1720.
39. Olofsson, S. O., Boström, P., Lagerstedt, J., Andersson, L., Adiels, M., Perman,
J. Rutberg, M., Li, L., and Borén, J. 2009. The lipid droplet: A dynamic organelle,
not only involved in the storage and turnover of lipids. Pages 1-26 in: Cellular
Lipid Metabolism. C. Ehnholm, ed. Springer-Verlag, Berlin.
40. Orion, D., and Wergin, W. P. 1982. Chloroplast differentiation in tomato root galls
induced by the root-knot nematode Meloidogyne incognita. J. Nematol. 14:77-83.
41. Pece, S., Tosoni, D., Confalonieri, S., Mazzarol, G., Vecchi, M., Ronzoni, S.,
Bernard, L., Viale, G., Pelicci, P. G., and Di Fiore, P. P. 2010. Biological and
molecular heterogeneity of breast cancers correlates with their cancer stem cell
content. Cell 140:62-73.
42. Schroeder, N. E., and MacGuidwin, A. E. 2007. Incorporation of a fluorescent
compound by live Heterodera glycines. J. Nematol. 39:43-49.
43. Sijmons, P. C., Grundler, F. M. W., von Mende, N., Burrows, P. R., and Wyss, U.
1991. Arabidopsis thaliana as a new model host for plant-parasitic nematodes.
Plant J. 1:245-254.
44. Sobczak, M., Golinowski, W., and Grundler, F. M. W. 1997. Changes in the
structure of Arabidopsis thaliana roots induced during development of males of
85
the plant parasitic nematode Heterodera schachtii. Eur. J. Plant Pathol. 103:113-
124.
45. Urwin, P. E., Møller, S. G., Lilley, C. J., McPherson, M. J., and Atkinson, H. J.
1997. Continual green-fluorescent protein monitoring of cauliflower mosaic virus
35S promoter activity in nematode-induced feeding cells in Arabidopsis thaliana.
Mol. Plant-Microbe Interact. 10:394-400.
46. Vercauteren, I., Van Der Schueren, E., Van Montagu, M., and Gheysen, G. 2001.
Arabidopsis thaliana genes expressed in the early compatible interaction with
root-knot nematodes. Mol. Plant-Microbe Interact. 14:288-299.
47. Vieira, P., Banora, M. Y., Castagnone-Sereno, P., Rosso, M. N., Engler, G., and
de Almeida-Engler, J. 2012. An immunocytochemical procedure for protein
localization in various nematode life stages combined with plant tissues using
methylacrylate-embedded specimens. Phytopathology 102:990-996.
48. Vieira, P., Danchin, E. G. J., Neveu, C., Crozat, C., Jaubert, S., Hussey, R. S.,
Engler, G., Abad, P., de Almeida-Engler, J., Castagnone-Sereno, P., and Rosso,
M. N. 2011. The plant apoplasm is an important recipient compartment for
nematode secreted proteins. J. Exp. Bot. 62:1241-1253.
49. Vieira, P., Engler, G., and de Almeida-Engler, J. 2012. Whole-mount confocal
imaging of nuclei in giant feeding cells induced by root-knot nematodes in
Arabidopsis. New Phytol. 195:488-496.
50. Wyss, U., and Grundler, F. M. W. 1992. Feeding behavior of sedentary plant
parasitic nematodes. Neth. J. Plant Pathol. 98 (Suppl.):165-173.
86
51. Wyss, U., Grundler, F. M. W., and Münch, A. 1992. The parasitic behavior of
second-stage juveniles of Meloidogyne incognita in roots of Arabidopsis thaliana.
Nematologica 38:98-111.
52. Xu, W., Ding, G., Yokawa, K., Baluska, F., Li, Q. F., Liu, Y., Shi, W., Liang, J.,
and Zhang, J. 2013. An improved agar-plate method for studying root growth and
response of Arabidopsis thaliana. Sci. Rep. 3:1273.
53. Yokawa, K., Kagenishi, T., Kawano, T., Mancuso, S., and Baluska, F. 2011.
Illumination of Arabidopsis roots induces immediate bursts of ROS production.
Plant Signal. Behav. 6:1460-1464.
54. Zunke, U. 1990. Observations on the invasion and endoparasitic behavior of the
root lesion nematode Pratylenchus penetrans. J. Nematol. 22:309-320.
55. Zurbriggen, M. D., Carrillo, N., Tognetti, V. B., Melzer, M., Peisker, M., Hause, B.,
and Hajirezaei, M. R. 2009. Chloroplast-generated reactive oxygen species play
a major role in localized cell death during the non-host interaction between
tobacco and Xanthomonas campestris pv. vesicatoria. Plant J. 60:962-973.
87
2. 8. FIGURES
Fig. 1. Arabidopsis grown in microscopy rhizosphere chambers (micro-ROC). For all experiments, a single Arabidopsis seedling was grown in each micro-ROC. The developing root system grows between a nylon mesh and glass slide but has access to the potting mix. The glass slide is covered with a black plastic film (removed here) to block out light. Nematode inoculum was pipetted between the nylon mesh and glass slide to maximize infection.
88
Fig. 2. Pratylenchus penetrans stained with PKH26. A, P. penetrans 3 days after labeling with PKH26 outside of plant, stored at 4°C until imaging. Intense fluorescence of intestinal lipids. All other pictures show nematodes in planta. B, P. penetrans at 3 days after inoculation (DAI) in Arabidopsis roots and C, at 19 DAI. D, Molting female at
19 DAI; arrow indicates metacorpus. E, Resting male at 26 DAI, arrow indicates spicules. F, Anterior portion of female during migration through epidermal tissue at 27
DAI. G, Migrating female at 27 DAI with deposited egg (arrow). H, Deposited eggs at 27
DAI (arrows). Scale bars = 50 µm.
89
90
Fig. 3. Heterodera schachtii stained with PKH26. A, H. schachtii second-stage juveniles
(J2s) 3 days after labeling with PKH26 outside of plant, stored at 4°C until imaging.
Intense fluorescence of intestinal lipids. All other pictures show nematodes in planta. B,
Intracellular migration of J2 at 10 days after inoculation (DAI). C, J3 at 14 DAI begins to break through root cortex. D, Developing J3 or J4 with median bulb (arrow) at 22 DAI.
E, Molting J3 at 23 DAI. Arrows indicate shed cuticle. F, Developing female at J4 stage,
23 DAI. G, Developing male at J4 stage, 23 DAI. H, Adult female during oogenesis
(arrow indicates developing eggs) at 23 DAI. I, Mature eggs (top arrow) and vulval cone
(bottom arrow) at 28 DAI. J, Syncytium (arrow) at 28 DAI. Scale bars = 50 µm.
91
92
Fig. 4. Meloidogyne chitwoodi stained with PKH26. A, M. chitwoodi second-stage juveniles (J2s) 3 days after labeling with PKH26 outside of plant, stored at 4°C until imaging. Intense fluorescence of intestinal lipids. All other pictures show nematodes in planta. B, Migrating J2 at 3 days after inoculation (DAI). C, Late parasitic J2 and developing gall at 12 DAI. D and E, J2 (arrow) at 12 DAI during intercellular migration.
F, Early giant cell formation at 12 DAI. G, J3 or J4 breaking through the root cortex at 15
DAI. H and I, Posterior end of female surrounded by gall tissue at 27 DAI with egg
(arrow). Scale bars = 50 µm.
93
94
Fig. 5. Peroxisomes tagged with yellow fluorescent protein (YFP) in transgenic
Arabidopsis roots infected with Meloidogyne chitwoodi. A, Distribution of YFP-tagged peroxisomes in uninfected roots 22 days after germination in microscopy rhizosphere chambers. B, Early M. chitwoodi gall formation at 3 days after inoculation (DAI). C,
Increased peroxisome concentration in developing gall region compared with surrounding uninfected root tissue at 8 DAI. D to G, Reduction of peroxisome concentration in developing gall region compared with surrounding uninfected root tissue at 12 DAI (D), 19 DAI (E), and 27 DAI (F and G). Scale bars = 50 µm.
95
Fig. 6. Quantification of yellow fluorescent protein-tagged peroxisomes in Meloidogyne chitwoodi-infected and uninfected Arabidopsis roots. A, Number of peroxisomes per
0.01 mm2 of root. B, Area of peroxisomes (1 × 10–3 mm2) per 0.01 mm2 of root.
96
CHAPTER THREE
MELOIDOGYNE INCOGNITA EFFECTOR 16D10 MODIFIES XYLEM
DIFFERENTIATION IN ARABIDOPSIS THALIANA TO
FACILITATE NEMATODE PARASITISM
Phuong T.Y. Dinh1, Lei Zhang1, Debra A. Inglis2, and Axel A. Elling1
1Department of Plant Pathology, Washington State University, Pullman, WA 99164
2Department of Plant Pathology, Washington State University, WSU Mount Vernon
Northwestern Washington Research & Extension Center, 16650 State Route 536,
Mount Vernon, WA 98273
Corresponding author : Axel A. Elling
E-mail: [email protected]
Following the format of the Molecular Plant-Microbe Interactions
97
3. 1. ABSTRACT
The root-knot nematode (RKN) Meloidogyne incognita is one of the most destructive plant parasitic nematodes and affects all major agricultural systems. This sedentary endoparasitic nematode species totally depends on host nutrients it absorbs from root feeding sites. Nematode effectors, proteinaceous secretions from the nematode esophageal gland cells, are essential for plant-nematode interactions at the molecular level, and they function in establishing parasitism by modifying plant growth, development and immunity during the formation of feeding sites in plant roots. In this study, the roles of 16D10 effector in M. incognita parasitism were investigated using
16D10-overexpressing Arabidopsis thaliana lines. The results showed that 16D10 increased plant susceptibility to M. incognita (increased the number of egg masses) but not to Pseudomonas syringae (no change in number of CFU mg-1 leaves), even though this effector suppressed the induction of the pathogen-related 5 gene in response to
P. syringae infection. In contrast, 16D10 enhanced callose deposition in response to a pathogen elicitor, flg22. Although no effect on root length was observed, 16D10 increased the number of undifferentiated metaxylem cells concurrently with induction of the gene expression of VND7, a marker for xylem development. Overall, the 16D10 effector played various roles in modifying the defense and development of A. thaliana to facilitate M. incognita parasitism.
98
3. 2. INTRODUCTION
Meloidogyne incognita is one of the major and virulent root-knot nematode (RKN) species and among the most damaging of soil-borne plant pathogens (Trudgill and Blok
2001). M. incognita threatens a wide range of crops worldwide, especially in tropical regions (Bird et al. 2009; Sasser 1977; Trudgill and Blok 2001). Enormous economic loss is caused by M. incognita due to the broad host range and widespread distribution of this species (Trudgill and Blok 2001). RKN derive host nutrients from feeding sites in the root, causing yield loss in crop production. Root galls are typical symptoms of RKN infection and are formed by the hypertrophy and hyperplasia of root cells surrounding
RKN and feeding sites (Niebel et al. 1996). Galls formed by RKN also cause quality loss in tuber/root crops such as potato, carrot and yam.
The life cycle of M. incognita starts with the first-stage juveniles (J1) in eggs. J1 molt into the second-stage juveniles (J2), which hatch from eggs. J2 migrate toward and invade host roots, where they become sedentary and induce the formation of five to eight giant cells per feeding site (Caillaud et al. 2008; Wyss et al. 1992). J2 molt into third and fourth-stage juveniles (J3 and J4). J4 develop into adults, either males or females. Because M. incognita are sedentary endoparasitic nematodes, the survival of
M. incognita is dependent on the maintenance of feeding sites during the entire molting period, from J2 to adult stages. Adult males leave host roots to fertilize females. Adult females remain sedentary and continue taking up host nutrients from giant cells. Mature females produce eggs in gelatinous egg matrices, called egg masses (Castagnone-
Sereno 2002; Castagnone-Sereno et al. 2007).
99
Giant cell formation is vital for M. incognita, and believed to be induced by RKN effectors. M. incognita effectors are defined as the proteins that are produced in the esophageal glands, one dorsal and two subventral. Effectors are injected into host cells by nematode stylets – hollow and needle-like structures (Davis et al. 2004; Hussey
1989; Hussey and Mims 1990) used for feeding. RKN effectors have been localized at the subcellular level in the apoplasm, cytoplasm and nucleus of affected plant cells
(Jaouannet et al. 2012; Rosso et al. 2011; Vieira et al. 2011; Zhang et al. 2014a).
M. incognita effectors suppress host defenses effectively. For example,
M. incognita calreticullin effector (Mi-CRT) was characterized functioning in suppression of PTI, pathogen-associated molecular pattern (PAMP)-triggered immunity (Jaouannet et al. 2013). Moreover, RKN effectors are thought to modify the division and fate of host cells inducing the differentiation of root cells for transformation into hypertrophied, metabolically hyperactive and multinucleated giant cells (Caillaud et al. 2008). The effector Mi8D05 from M. incognita is the case of changing host plant growth, e.g., increasing the height of A. thaliana; while Mi8D05 was found to facilitate solute and water transport in giant cells during the course of M. incognita parasitism (Xue et al.
2013).
Numerous studies have examined the M. incognita 16D10 effector. Huang and colleagues (2006b) found that, in contrast to Mi8D05, 16D10 did not alter shoot development but increased root length. 16D10 cloned from M. incognita was highly conserved in Meloidogyne species, including M. arenaria, M. javanica, M. hapla and
M. chitwoodi (Dinh et al. 2014a; Huang et al. 2006b). 16D10 plays an essential role in the parasitism of RKN because the silencing of 16D10 by RNA interference in planta
100 significantly reduced the reproduction of Meloidogyne species in both A. thaliana and potato (Dinh et al. 2014a; Dinh et al. 2014b; Huang et al. 2006a). Furthermore, overexpression of 16D10 in tobacco hairy roots increased root length and stimulated lateral root formation (Huang et al. 2006b). Overexpressed 16D10 A. thaliana plants had longer primary roots than control plants (Huang et al. 2006b). The mechanism of
16D10 in modifying root growth is not clear although 16D10 may interact with
SCARECROW, a transcription factor (Huang et al. 2006b).
In M. incognita, full-length16D10 consists of 43 amino acids (aa) including 30 aa in a signal peptide and 13 aa in a mature peptide (Huang et al. 2006b). The amino acid sequence of 16D10 mature peptide is similar to the CLE motif of the B-type CLE
(CLAVATA3/ENDOSPERM SURROUNDING REGION) in A. thaliana (Mitchum et al.
2008). One of the B-type CLE peptides, CLE41, promoted vascular division but suppressed xylem differentiation (Etchells and Turner 2010; Whitford et al. 2008). The
CLE-like effectors of cyst nematodes terminated shoot apical meristem, decreased root length and complemented the phenotype of clv3 mutant (Lu et al. 2009; Wang et al.
2011; Wang et al. 2005). However, overexpression of 16D10 was not able to restore function lost in the clv3 mutant (Huang et al. 2006b).
Another study demonstrated CLE41 involvement in the ethylene pathway of plant defenses (Etchells et al. 2012). An interaction between PHLOEM INTERCALATED
WITH XYLEM (PXY, a CLE41 receptor) and ERECTA (ER, a leucine-rich repeat receptor-like kinase) regulated the organization of A. thaliana vascular tissue (Etchells et al. 2013). In addition, ERECTA played a role in plant defense mechanisms (Godiard et al. 2003). Considering this, the hypothesis of this study is that similar to CLE41,
101
16D10 increases root cell division and suppresses host defense to facilitate M. incognita parasitism.
3. 3. MATERIALS AND METHODS
Subcellular localization
M. incognita 16D10 mature peptide (DQ087264) was fused with enhanced green fluorescent protein (eGFP) and β-glucuronidase (GUS) following the procedure published by Zhang and colleagues (2014a). In detail, the sequence of the 13 aa of
16D10 mature peptide was cloned into pGUS-ENTR-3 and pGUS-ENTR-5 vectors before being transferred into destination vectors, pSITE-2CA and pSITE-2NB, respectively (Chakrabarty et al. 2007; Zhang et al. 2014a). Primers for the cloning,
16D10-EcoRI, 16D10-BamHI-1 and 16D10-BamHI-2, are shown in Table S1. After cloning, eGFP and GUS were fused at the C-terminal or N-terminal of 16D10 mature peptide. These final vectors, carrying the eGFP-GUS-16D10 or 16D10-GUS-eGFP under 35S promoter, were imported into Agrobacterium tumefaciens GV3101 by electroporation. An empty vector served as a control.
A. tumefaciens strains carrying either eGFP-GUS-16D10 or 16D10-GUS-eGFP were separately co-infiltrated with a suppressor of post-transcriptional gene silencing,
HcPro of Potato virus Y (Brigneti et al. 1998). To do that, each strain of A. tumefaciens was suspended in an infiltration buffer, [10 mM 2-(N-morpholino) ethanesulphonic acid
(MES) at pH 5.6, 10 mM MgCl2 and 150 µM acetosyringone], to obtain OD600 (optical density at wavelength 600 nm) of 1 (Bendahmane et al. 2000). The bacterial suspensions were incubated at 25ºC for 2 h. A mixture of eGFP-GUS-16D10 or 16D10-
102
GUS-eGFP and HcPro was co-infiltrated into four leaves of two 4-week-old
N. benthamiana plants. The localization of 16D10 was detected at three days after infiltration by observing the eGFP fluorescent signal, under an LSM 510 META inverted confocal microscope (Carl Zeiss. Jena, Germany: excitation/emission wavelengths of
488/510-550 nm).
Bimolecular fluorescence complementation (BiFC)
The results of the Y2H assay (Fig. S1) on the interactions of plant proteins and
16D10 were confirmed in planta by BiFC (Martin et al. 2009). In brief, mature 16D10 peptide was expressed from the pSITE-BiFC-C1, and the A. thaliana target proteins
(identified by Y2H) were cloned in the pSITE-BiFC-N1 vectors, to construct pSITE-
BiFC-C1-16D10 and pSITE-BiFC-N1-target, respectively. Primers for the cloning,
16D10-GWF1, 16D10-GWR1, 16D10-GWR2, NTF-GWF, NTF-GWR, CF-GWF, CF-
GWR, PLC2-GWF, PLC2-GWR, PCK-GWF and PCK-GWR, are shown in Table S1.
These fusion vectors were imported into A. tumefaciens GV3101 by electroporation.
The A. tumefaciens suspensions for infiltration were prepared as described above, except that the OD600 values of these suspensions were 1.8. For BiFC, a mixture of equal ratio of the HcPro, pSITE-BiFC-C1-16D10, and each pSITE-BiFC-N1-target was co-infiltrated into N. benthamiana leaves (two sites per leaf, three leaves per plant and two plants for each mixture). These infiltrated leaves were harvested at three days after infiltration and observed under the LSM 510 META inverted confocal microscope. The signals of enhanced yellow fluorescent protein (eYFP) that were detected in
N. benthamiana leaves indicated the interaction of 16D10 and A. thaliana target proteins.
103
Overexpression of 16D10 in A. thaliana
The full-length sequence and the last 39 nucleotide sequence of 16D10 open reading frame (ORF) were cloned into pBI121 vector (CD3-388) to overexpress the full- length or mature 16D10 peptides, respectively, under the control of the 35S promoter.
The sequences of primers, 16D10-BIF1, 16D10-BIF2 and 16D10-BIR are shown in
Table S1. The sequences of these two overexpression constructs were confirmed by sequencing at Elim Biopharmaceuticals (Hayward, CA, USA). These two vectors were introduced into A. tumefaciens strain GV3101 by electroporation; the overexpression constructs were incorporated into the genome of A. thaliana ecotype Col-0 using the floral dip method (Clough and Bent 1998). Transgenic A. thaliana seeds were screened for the insertion of the 16D10 transgene by germinating them on selective media, one- half strength Murashige & Skoog basal salt media – MS media (Caisson Labs, North
Logan, UT), supplemented with 1.5% D-sucrose, 0.6% Daishin agar, timentin at 0.5 g/liter, and kanamycin (Plant Media, Dublin, OH) at 50 µg/ml. After being screened, four lines were chosen: two lines with the mature 16D10 expression (without the signal peptide), named OX16D10-1 and OX16D10-2, and the other two lines, OX-16D10SP-1 and OX-16D10SP-3, having the full 16D10 peptide (with the signal peptide) overexpressed. One hundred percent of the third-generation (T3) of transgenic seeds germinated on the selective media meaning the T3 seeds were homologous for the
16D10 transgenes. The T3 seeds of each line were used as plant material in further experiments/assays.
104
Plant growth conditions
A. thaliana seeds were surface sterilized for 5 min in 70% ethanol and then 5 min in 2.5% NaOCl, followed by being washed 10 times with sterile water. The sterilized seeds were plated individually on one-half strength MS media plus 1.5% D-sucrose and
0.6% Daishin agar. Seeds were vernalized at 4ºC for two to four days before germinating in a growth chamber (22ºC with 12 h photoperiod). After germination, the seedlings were transferred to various media/potting mix to conduct various experiments, described below. The four overexpression lines, OX16D10-1, OX16D10-2,
OX16D10SP-1 and OX16D10SP-3, were used as plant materials for further experiments to compare with control lines, wild type Col-0 (named as COL in this study) and the 16D10 silencing line (D4) generated previously (Dinh et al. 2014a).
For the M. incognita infection assay, the seedlings of the six A. thaliana lines were transferred to infection media, Gamborg’s B-5 media (Caisson Labs, North Logan,
UT) plus 0.6% Daishin agar at three days after germination (DAG). Three plants per square plate (Fisher Scientific, Fair Lawn, NJ), six plates per A. thaliana line were grown under the same growth chamber conditions for another 14 days before being inoculated with M. incognita J2. The plates were placed in a vertical position on growth chamber racks to allow A. thaliana root growth along the surface of media.
For P. syringae DC3000 infection and callose deposition assays, 7 DAG seedlings of six A. thaliana lines were transferred into moistened Sunshine No. 1 potting mix (Sun Gro Horticulture, Agawam, MA). These seedlings were grown in a growth chamber at 22ºC with 16 h daylight and at 20ºC with 8 h no light for another 21 days.
The seedlings were watered every two days with 20-10-20 NPK liquid fertilizer.
105
In order to analyze the root length and root anatomy of A. thaliana lines, 3 DAG, six A. thaliana lines (wild type COL, OX16D10-1, OX16D10-2, OX16D10SP-1,
OX16D10SP-3 and D4) were transferred to the same plate of one-half strength MS media supplemented with 1.5 % D-sucrose and 0.6% Daishin agar (one plant per line).
Twelve plates per experimental replication were grown in the growth chamber for seven more days. The plates again were positioned vertically. Root lengths were measured by
ImageJ (written by Wayne Rasband at the National Institute of Health, Bethesda, MD,
USA) after the plates were scanned by Epson perfection V600 at 10 DAG. Following, 2 mm root segments (5 mm above root tips) of all experimental lines were harvested for the immuno-labelling of A. thaliana xylem. The root systems of all studied A. thaliana lines also were harvested at 10 DAG for analyzing gene expressions. The experiment was repeated three times.
Nematode inoculum
M. incognita culture was maintained on Solanum lycopersicum ‘Rutgers’ grown in autoclaved sand under greenhouse conditions (16 h daylight at 24ºC and 8 h nighttime at 20ºC). M. incognita eggs were extracted from infested tomato roots (around 3 months after inoculation) using 0.5% NaOCl (Hussey and Barker 1973). After 3 min agitation in
0.5% NaOCl, the root suspension was poured over a nested stack of test sieves (850,
75, and 25 µm pore sizes from top to bottom). Eggs were collected from the finest sieve
(25 µm pore size) after being washed with tap water for 2 min. Eggs then were purified on a sucrose gradient by mixing 20 ml egg suspension and 20 ml of 70% sucrose in a
50 ml tube. Water was gently added to the top layer of the tube and the tube was then spun at 375 x g for 3 min in a clinical centrifuge. After that, eggs at the interface of the
106 sucrose gradient were harvested and rinsed for about 10 min on the sieve with the finest mesh. Three days after the eggs hatched at 24ºC, infective J2 were harvested from them.
The surface sterilization of infective J2 was conducted following the protocol from
Iberkleid and colleagues (2013) with some modifications. J2 were placed on a sterile
FMTM-Millipore-Membrane filter (SSWP002500) in a sterile Millipore Swinnex holder
(SX0002500, Merck, Darmstadt, Germany). Ten milliliters of 0.01% (w/v) mercuric chloride (HgCl2) (Acros Organics, Fair Lawn, NJ) and 10 ml of 0.7% Streptomycin solution (Research Products International, Corp., Mount Prospect, IL) were consecutively filtered through the membrane. After that, 100 ml of sterilized water were also passed through the membrane. J2 on the membrane were re-suspended in sterilized water to a final concentration of 2500 J2 per ml. The J2 suspension was used as inoculum for the infection assay.
M. incognita infection assay
The 17 day-old A. thaliana plants on Gamborg’s media (described above) were inoculated with the sterilized M. incognita J2 (250 J2/plant, 3 plants/plate and 6 plates/line). The inoculated plants were vertically grown in the growth chamber. At 35 days after inoculation (DAI), each plate was filled with 40 ml of phloxine B (Fisher
Scientific) solution at 0.15 g/liter for 18 h at 24ºC to stain egg masses produced by
M. incognita. The phloxine B solution was replaced by water for two days in order to remove the nonspecific phloxine B stains. After that, the plates were scanned by the
Epson perfection V600 scanner and the egg masses were counted under a Stemi
107
2000C stereomicroscope (Zeiss) at 20x magnification. The experiment was repeated three times; each time had six replicates (six plates) per line.
Pseudomonas syringae DC3000 infection
Following a protocol published by Zhang and colleagues (2014b), three leaves of each 28 day-old A. thaliana seedling in potting mix were infiltrated with 105 P. syringae
DC3000 cells in 10 mM MgCl2. For studying gene expression, 20 h after infiltration, about 50 mg leaves of all studied lines were harvested and frozen at -80ºC. To assess the susceptibility of the A. thaliana lines to P. syringae, the leaves of four seedlings (in four replicates) were harvested for each line at each time point, 0 DAI and 3 DAI. For the control treatment, A. thaliana leaves were infiltrated with 10 mM MgCl2 only and harvested at 3 DAI. For each seedling, harvested leaves were weighed and ground in 1 ml of 10 mM MgCl2. P. syringae titers (CFU/g leaf fresh weight) were determined by serial dilutions and plating on King’s medium B [2% proteose peptone #3 (BD Difco),
0.15% K2HPO4, 1% glycerol, 6 mM MgSO4 and 1.5% agar with 50 mg/l of rifampicin and
50 mg/l ampicillin]. P. syringae infection assays were repeated twice and representative data are shown in the results.
Callose depositions
28 day-old A. thaliana seedlings in potting mix were used as plant material for callose deposition evaluations following a protocol modified from one described by Kim and colleagues (2005). The whole leaf of each seedling was infiltrated with 1 µM flagellin22 (flg22, PhytoTechnology Laboratories, Shawnee Mission, KS) dissolved in sterilized water. In the same seedling, another leaf was infiltrated with sterile water as a control. The assay was repeated three times and each time involved six lines (wild type
108
COL, OX16D10-1, OX16D10-2, OX16D10SP-1, OX16D10SP-3 and D4) and six seedlings per line. The infiltrated leaves were harvested at 24 h after infiltration and were fixed in 95% ethanol immediately after harvesting. The fixation step lasted at least
24 h with one or two exchanges of fresh 95% ethanol until the leaves were translucent.
These leaves were gradually rehydrated using ethanol solutions of 70% and 50% for 2 h each, and finally by sterile water for 16 h. Then, the leaves were incubated in phosphate buffer (0.07 M Na2HPO4, pH 9) for 2 h before being stained for 16 h at 4ºC in 0.01% aniline blue dissolved in phosphate buffer. Callose deposition was visualized under an
AxioObserver A1 inverted microscope equipped with a mercury lamp (X-cite Series 120, and a DAPI filter set (excitation at 365 nm and emission ranging from 445 to 450 nm wavelength) at 50x magnification. The callose depositions of six different areas (avoid from the infiltrated positions) in each leaf were captured and quantified with ImageJ following previously established procedures (Desai and Hu 2008). Briefly, RGB color images were transformed to 8-bit black-and-white images. The “threshold” function was adjusted to designate black pixels as callose depositions. The number of callose depositions in each image was counted by the “analyze particles” function of ImageJ.
Immuno-labelling of A. thaliana xylem
The immuno-labelling was conducted following the procedure described by
Davies and colleagues (2012) with some modifications. In detail, the 2 mm root segments that were harvested at 5 mm above the root tips from 10 day-old A. thaliana seedlings were fixed immediately in 4% formaldehyde (Ted Pella, INC, Redding, CA) mixed in 0.2 M phosphate buffer, pH 7 at 4ºC for 16 h. The formaldehyde was washed away with phosphate-buffered saline (PBS) buffer three times. After that, these root
109 samples were dehydrated consecutively with an ethanol series of 10, 20, 30 and 50% for 15 min at each concentration. The dehydration of the root samples continued with a subsequent ethanol series of 70, 90 and then 100% for 30 min at each concentration.
The dehydrated root samples were embedded using a step-wise resin series of 10, 20,
30, and 50% of LR white acrylic resin (diluted with 100% ethanol) for 30 min at each step. Higher concentrations (70, 90 and 100%) of the resin were consecutively exchanged for 1 h each. Root samples were embedded by the replacement of fresh LR white acrylic resin (100%) three times in two days before being molded in gelatin capsules over 24 h at 55ºC.
Transverse sections (1 µm thickness) of root samples were sliced with a Leica
Reichert Ultracut R microtome and mounted onto PTFE-coated eight-well glass slides
(MP Biomedicals). The root sections on the slides were blocked with 5% (w/v) milk protein in PBS buffer for 30 min before being incubated for 2 h at 37ºC in LM11 primary antibodies (PlantProbes, UK) diluted five times in PBS. Unbound LM11 antibodies were carefully rinsed off three times with PBS, followed by incubation for 1.5 h in the dark with anti-rat immunoglobulin G conjugated to fluorescein isothiocyanate (FITC, Sigma,
St. Louis, MO) diluted 100 fold in PBS. The root sections were rinsed again with PBS three times before being stained in the dark with 1 mg/ml Calcoflour-white (Fluka,
Sigma). Finally, the slides were rinsed thoroughly with PBS three times and mounted in glycerol-based Citifluor AF1 antifade (Electron Microscopy Sciences, Hatfield, PA). The root sections were captured with the LSM 510 META inverted confocal microscope. The microscope multi-tracking was adjusted with two sets of excitation/emission
110 wavelengths, 405/500-530 nm (in blue indicating the fluorescent signal of Calcoflour- white) and 488/530 nm (in green indicating the fluorescent signal of FITC).
Study of gene expression with quantitative real-time polymerase chain reaction
(qRT-PCR)
To study the levels of gene expressions, total RNA of A. thaliana samples were extracted following the instructions of the RNeasy Plant Mini kit (Qiagen, Valencia, CA).
A. thaliana cDNA of each sample was synthesized from 1 µg of the total RNA with iScriptTM cDNA Synthesis (Bio-Rad, Hercules, CA). The expression levels of xylem marker genes and genes encoding for pathogen-related (PR) proteins were compared among six A. thaliana lines. The common xylem marker genes are VASCULAR-
RELATED NAC-DOMAIN genes (VND6 and VND7) as well as IRREGULAR XYLEM genes (IRX1, IRX3 and IRX5). The expression levels of PR genes (PR1, PR2 and PR5) also were analyzed with qRT-PCR.
qRT-PCR was performed in a CFX96 Real-Time PCR Detection System (Bio-
Rad) with SsoAdvancedTM Universal SYBR Green Supermix (Bio-rad). The ubiquitin carboxyl-terminal hydrolase 22 (UBP22) was chosen as an internal control gene for qRT-PCR as recommended for M. incognita-infected root tissues by Hofmann and
Grundler (2007). Primers for qRT-PCR are shown with suffix “RTF” and “RTR” in Table
S1. For each A. thaliana line, three technical replicates of three root samples (biological replicates) were run with the program: 95ºC for 3 min; 39 cycles at 95ºC for 10 s; and
60ºC for 1 min. The procedure was followed by a melt-curve analysis with a temperature increase of 0.5ºC for 5 s after each cycle from 65ºC to 95ºC. The relative gene
111 expressions of the xylem marker genes were analyzed by the CFX Manager 3.1 program (Bio-Rad).
Data analysis.
All data were analyzed with Microsoft Excel (to calculate means and standard errors as well as to generate bar graphs), and with SAS 9.2 software (to determine statistical significance using a Student’s t test at α level = 0.05). Subsequent image processing was performed with ImageJ and Adobe Photoshop.
3. 4. RESULTS
The localization and interaction of effector 16D10 in N. benthamiana
To investigate the localization of a target protein in N. benthamiana cells, the protein is fused with both enhanced green fluorescent protein (eGFP) and β- glucuronidase (GUS), which helps to prevent the passive diffusion of the target protein among subcellular compartments (Grebenok et al. 1997; Haasen et al. 1999; Zhang et al. 2014). Therefore, the fusion of 16D10 mature peptide with both GUS and eGFP allowed 16D10 to be localized as green fluorescent signals in the cytoplasm of any tobacco cells carrying eGFP-GUS-16D10 (Fig. 1A) as well as 16D10-GUS-eGFP (Fig.
1B).
Screening with the A. thaliana cDNA library, Y2H indicated the interaction of
16D10 with four plant proteins: nuclear transport factor 2 (NTF2, AT5G60980); cytosolic factor (CF, AT1G72150); phosphoinositide-specific phospholipase C2 (PLC2,
AT3G08510); and pfkB-like carbohydrate (PCK, AT1G19600). These interactions were validated by the co-transformation of 16D10 mature peptide and the individual target
112 proteins in yeast cells (Fig. S1). Yeast colonies that were able to grow on the quadruple dropout media indicated that 16D10 mature peptide interacted with the four target proteins (Fig. S1).
After the interactions between 16D10 and the target proteins were validated in yeast system, they were confirmed in N. benthamiana cells with the BiFC (Fig. 1C to F).
The CF and PLC2 bound with 16D10 to recover the eYFP signals in the cytoplasm (Fig.
1C and D). The complex of 16D10-PCK was localized in both cytoplasm and nucleus
(Fig. 1E). The eYFP signal from the 16D10-NTF2 complex was detected in the intercellular spaces of the tobacco cells (Fig. 1F).
Overexpression of 16D10 increased the susceptibility of A. thaliana to
M. incognita but not to P. syringae
Overexpression of 16D10 full-length under 35S promoter in A. thaliana created
OX16D10SP-1 and -3 lines; overexpression of 16D10 encoding mature peptide created
OX16D10-1 and -2 lines. The expression of 16D10 transgene in transgenic lines was confirmed by RT-PCR (Fig. S2). The OX16D10-1 had the highest 16D10 expression level among the other transgenic lines while the expression of 16D10 was not detected in wild type COL. The RT-PCR also detected the RNAi of 16D10 silencing line, D4 (Fig.
S2).
Overexpression of 16D10 (either full-length or mature peptides) significantly increased the numbers of M. incognita egg masses (stained with phloxine B in pink) produced in the transgenic A. thaliana (Fig. 2A, B and C) (P < 0.05). At 35 DAI, even though M. incognita produced fewer egg masses in OX16D10SP-3 (34.44) than in
OX16D10-1 (45.89) (P < 0.05), the average number of egg masses per plant in the
113
OX16D10SP-3 still was significantly higher than COL control (21.5). The egg mass numbers in OX16D10-2 (45.22) and OX16D10SP-1 (39.33) also were higher than in
COL, but not significantly different than in the other 16D10-overexpressing lines (Fig.
2C). D4, a 16D10 silencing line, had the lowest number of egg masses (11.39) (P <
0.05).
Although, overexpression of 16D10 increased the susceptibility of A. thaliana to
M. incognita in this study (Fig. 2C), there was no statistically significant difference in
P. syringae susceptibility among any of the investigated lines (Fig. 3). Both of COL and
D4 control lines as well as 16D10-overexpressing lines had similar levels of the infiltrated P. syringae inside the A. thaliana leaves (day 0, data not shown). At three days after infiltration, there was no contamination in the control leaves infiltrated with 10 mM MgCl2. The log10 values of P. syringae CFU/g of fresh leaves were not different in any of the A. thaliana lines (from 4.97 to 5.09; P > 0.05) at three days after infiltration.
The effects of overexpression of 16D10 in the basal defense of A. thaliana
OX16D10-1 leaves had more callose deposition than leaves of COL wild type in response to flg22 (Fig. 4A and B). The leaves of those lines that were infiltrated with sterile water showed no callose deposition (Fig. 4C and D). In contrast, the leaves of all
16D10-overexpressing lines had significantly higher levels (P < 0.05, Fig. 4E) of callose deposition in response to flg22 than did leaves of COL (282.32 per mm2) and D4 (296.1 per mm2). Among the transgenic lines, OX16D10SP-3 had the highest number of callose depositions (613.01 per mm2). The numbers of callose depositions per mm2 in the other transgenic lines, OX16D10-1 (510.64), OX16D10-2 (443.32) and
114
OX16D10SP-1 (507.96), did not differ significantly (P > 0.05); all values were significantly lower than in OX16D10SP-3 (P < 0.05).
Overexpression of 16D10 prevented the induction of PR5, a gene involved in plant basal defenses in response to 20 h infiltration with P. syringae (Fig. 5). When the expression of PR5 in COL leaves was set at 1 as control, the relative expression of PR5 in 16D10-overexpressing lines, OX16D10-1, -2, OX16D10SP-1, and -3 were 0.52, 0.23,
0.36 and 0.3 respectively (Fig. 5). Therefore, the induction of PR5 gene in response to
P. syringae was reduced about two to four fold in 16D10-overexpressing lines compared to COL. In contrast to the overexpressed lines, D4 had similar induction of
PR5 gene compared to COL (P > 0.05). The induction of other PR genes (PR-1 and
PR-2) was not changed by the overexpression of 16D10 (data not shown).
Overexpression of 16D10 did not change A. thaliana root lengths but increased the number of metaxylem cells in A. thaliana vascular bundles
Overexpression of 16D10, both the full-length and mature peptides, did not change the A. thaliana overall phenotype, including root lengths (Fig. 6A). The average root lengths of A. thaliana lines, COL, OX16D10-1 and -2, OX16D10SP-1 and -3 as well as D4 ranged from around 45 to 55 mm at 10 DAG. The difference in the average root lengths among these A. thaliana lines was not statistically significant (P > 0.05).
Even though 16D10 did not affect A. thaliana root elongation, this effector might have increased root cell division. Immuno-labelling the root cross-sections (about 5 mm above the root tips) of A. thaliana lines with LM11 enabled the visualization of the mature metaxylem cells. LM11 specifically bound to the mature metaxylem cells and also to the secondary antibody conjugated with FITC that had fluorescent signals
115
(shown as white asterisks in Fig. 2B to G). In the root cross-sections, there were undifferentiated metaxylem cells (shown as white arrowheads) in between the two mature metaxylem cells. The numbers of the mature metaxylem cells of all investigated
A. thaliana lines were not different (two cells in each cross-section), but 16D10 overexpressing lines had more undifferentiated metaxylem cells (two cells in Fig. 6C to
F) than wild type COL and D4 (one cell in Fig. 6B and G).
Overexpression of 16D10 changed the expression of A. thaliana genes involved in xylem differentiation
Because 16D10 transgenic lines had more metaxylem cells than did COL, the expressions of marker genes for xylem differentiation, such as VND6, VND7, IRX1,
IRX2 and IRX5, were analyzed in these lines. In 16D10 transgenic lines, only the expression level of VND7 was significantly higher than in COL (P < 0.05) (Fig. 7), while the expressions of the other genes (VND6, IRX1, IRX2 and IRX5) did not consistently differ among the lines except for OX16D10SP-3 (data not shown). When the expression level of VND7 in COL was set at 1 as control, the VND7 expressions in OX16D10-1,
OX16D10-2 and OX16D10SP-1 were nearly three times higher than in COL.
OX16D10SP-3 had the highest VND7 expression level, which was around six times higher than COL and twice the expression of the other transgenic lines. In contrast, the
VND7 expressions in D4 and COL did not differ.
116
3. 5. DISCUSSION
In this work, the various roles of 16D10 effector in modifying host cells were investigated using A. thaliana inoculated with M. incognita as a model system. 16D10 mature peptide (13 aa in length) localized in N. benthamiana cytoplasm. Previous studies have shown that the majority of M. incognita effectors (6F06, 2E07, 7A01, 7E12,
30G11, 5C03B, 2G02, 8H11, 1D08B, 6G07, 17H02 and 31H06) also localize in
N. benthamiana cytoplasm with the exception of 7H08 and Mi-EFF1; those effectors localize in the nuclear compartment where they are suspected to show transcriptional activation activity (Jaouannet et al. 2012; Zhang et al. 2014a).
In addition to plant cell cytoplasm and nucleus, the apoplasm is known to be another compartment targeted by RKN effectors, including cell wall modifying enzymes and Mi-CRT (Jaubert et al. 2005; Vieira et al. 2011). Nematode effectors, such as 7H08 and Mi-EFF1 from RKN as well as CLE-like effectors, Cellulose Binding Proteins (Hs
CBP) and ubiquitin extension protein from cyst nematodes, are transported from cytoplasm to apoplasm and/or nucleus (Jaouannet et al. 2012; Jaouannet and Rosso
2013; Zhang et al. 2014a). Effectors that carry nuclear localization signal/domains
(4E02, 6E07, Mi-EFF1 and 7H08) can actively traffic from cytoplasm to nucleus of host cells (Elling et al. 2007; Jaouannet et al. 2012; Zhang et al. 2014a). Some effectors require an interaction with a host protein in order to be transported to nucleus or apoplasm. Even though Hs CBP was found to interact with Arabidopsis pectin methylesterase protein 3 (PME3), there is no direct evidence showing that PME3 helps to transport Hs CBP from cytoplasm to apoplasm (Hewezi et al. 2008). The
117 translocation of 10A07, a cyst nematode effector, from cytoplasm to nucleus requires plant kinase to be phosphorylated (Hewezi et al. 2015).
In this study four A. thaliana target proteins, i.e., nuclear transport factor 2, cytosolic factor, phosphoinositide specific phospholipase C2 and pfkB-like carbohydrate, were found to interact with 16D10. The interactions demonstrated transport of 16D10 from cytoplasm to other plant cell compartments including nucleus and intracellular spaces. The complex of 16D10-NTF2 localized in intracellular spaces of tobacco cells while 16D10-PCK localized only in the nucleus of tobacco cells. 16D10 might function in a similar manner as cyst nematode effectors that traffic among cell compartments. Findings involving both Y2H and BiFC have revealed that 16D10 binds to multiple target proteins. These genes are involved in different development pathways of A. thaliana; thus, 16D10 might be a “sticky/promiscuous” protein, which can interact with multiple targets. This observation is in agreement with a study of Büttner and
Bonas (2002), in which an effector of Xanthomonas campestris pv. vesicatoria, HrpF, also was called “sticky” protein. Similar to this work with 16D10, investigations with Y2H could not identify the target protein that interacted with Mi-CRT (Jaouannet et al. 2013).
Like other nematode effectors that are secreted into host cells, 16D10 facilitates
M. incognita infection. Recent studies have found that the reproduction of RKN dramatically decreases if 16D10 is silenced by RNAi (Dinh et al. 2014a, 2014b, 2015;
Huang et al. 2006a). The importance of 16D10 effector in RKN infection was confirmed in this work. 16D10-overexpressing lines had about twice the number of M. incognita egg masses compared to COL control line. Also, M. incognita was more successful in parasitizing and reproducing in 16D10-overexpressing A. thaliana roots compared to the
118 control COL wild type. The effect of 16D10 on M. incognita reproduction is consistent with the effect of 7E12, Mi8D05 and calreticulin (Mi-CRT) effectors (Jaouannet et al.
2013; Souza et al. 2011, Xue et al. 2013). In those previous studies, overexpression of
7E12, Mi8D05 and Mi-CRT in A. thaliana facilitated gall formation by M. incognita so that gall numbers in the overexpressing lines were higher than in the control lines.
Mi-CRT effector has been reported to increase the susceptibility of A. thaliana to another root pathogen, Phytophthora parasitica (Jaouannet et al. 2013). Also, overexpression of 10A06, a cyst nematode effector, has been proven to increase the susceptibility of A. thaliana to multiple pathogens, including Heterodera schachtii,
Cucumber mosaic virus and P. syringae DC3000 (Hewezi et al. 2010). Even so, the susceptibility of 16D10-overexpressing A. thaliana lines to P. syringae DC3000 did not change in this study.
Mi-CRT and 10A06 effectors have been suggested to eliminate the innate immunity of A. thaliana thereby facilitating infection by multiple pathogens. In a report by
Jaouannet and colleagues (2013), callose depositions and induction of defense marker genes (PDF1.2, CYP81F2, WRKY29 and PAD4) in response to pathogen elicitor elf18 were suppressed by overexpression of Mi-CRT. However, the induction of the PR1 gene was not affected. The suppression of certain defense responses provides clear evidence of the ability of Mi-CRT to suppress host immunity (Jaouannet et al. 2013). As another example, 10A06-overexpressing A. thaliana lines that were inoculated with
H. schachtii had significantly lower expression levels of PR1, PR2 and PR5 (salicylic acid responsive genes) than control plants (Hewezi et al. 2010). Thus, 10A06 was
119 proposed to suppress salicylic acid signaling in order to increase the susceptibility of
A. thaliana to multiple pathogens (Hewezi et al. 2010).
In contrast to the case of Mi-CRT, overexpression of 16D10 in this work increased callose depositions in response to pathogen elicitor flg22. Callose depositions were reported to accumulate along the neighboring cell walls surrounding the giant cells during the early stage of giant cell formation (Hofmann et al. 2010). This deposition of callose is suggested to have a role in controlling the passive transport of water and solutes between giant cells and surrounding root cells (Hofmann et al. 2010). Therefore, the result of increased callose depositions in 16D10-overexpressing lines may suggest that the 16D10 effector plays a role in the enhancement of cell wall rigidity by increasing callose depositions during the early stage of giant cell formation.
Similar to the case of Mi-CRT and 10A06, 16D10 effector altered the expression of A. thaliana defense marker gene, PR5, in this study. 16D10 suppressed the induction of PR5 gene in response to P. syringae, although it had no effect on the inductions of the PR1 and PR2 genes. Even though the induction of PR5 was repressed, susceptibility to P. syringae in 16D10-overexpressing lines did not change. Therefore, the repression of PR5 induction by 16D10 may not be sufficient to alter the susceptibility of A. thaliana to P. syringae. However, the repression of PR5 might indicate the role of
16D10 effector in interrupting the salicylic acid signaling of A. thaliana.
To facilitate nematode parasitism, nematode effectors suppress host defenses and modify the developments and divisions host cells, particularly vascular parenchyma cells. Giant cells are surrounded by an extensive network of abnormal xylem cells that supply water and nutrients for giant cells (Jones and Dropkin 1976). The abnormal
120 xylem cells are thought to originate from parenchyma cells even though the mechanisms for formation and differentiation still are unknown. In this study, 16D10 effector was found to play a role in increasing the number of undifferentiated metaxylem cells of A. thaliana and in facilitating M. incognita infection. However, 16D10 did not change A. thaliana root length in this study.
Overexpression of 16D10 changed the expression level of VND7, but VND6 expression level was not changed. VND6 and VND7 are xylem marker genes and involved in the differentiation of xylem cells (Kubo et al. 2005; Yamaguchi et al. 2008).
When VND7 was overexpressed in A. thaliana, non-vascular cells were transdifferentiated into xylem vessel elements (Yamaguchi et al. 2008). Therefore, as suggested by this finding, the increase of VND7 expression might be responsible for the increase in number of undifferentiated xylem cells in the 16D10-overexpressing lines.
Hence, 16D10 effector potentially is involved in the induction of xylem formation/differentiation.
RKN are known to reprogram the cell cycle of the procambial/undifferentiated xylem cells in the vascular system of host roots to form giant cells (Williamson and
Hussey 1996; Wyss et al. 1992). Thus, it is interesting to hypothesize that RKN secrete effectors to modify vascular cells (including parenchyma, protoxylem and metaxylem), thereby enabling putative initial cells of giant cells to form (Bird 1961; Niebel et al.
1993). Indeed, Doyle and Lambert (2003) reported that overexpression of chorismate mutase (MjCM-1, a M. javanica effector) in A. thaliana inhibited vascular differentiation.
Soybean hairy roots with MjCM-1 overexpression failed to form lateral roots from lateral root meristems because the meristems did not generate vascular tissue. These authors
121 hypothesized that MjCM-1 effector altered plant cell development to facilitate
M. javanica infection. Increasing the number of undifferentiated metaxylem cells by overexpression of 16D10 might accelerate the formation of M. incognita feeding sites.
Accelerating feeding site formation by 16D10 could facilitate M. incognita parasitism in overexpressing A. thaliana lines.
Overall, similar to other reported M. incognita effectors (as reviewed by Hassan et al. 2010), 16D10 can play essential roles in the modification of the development and defense of host cells to facilitate RKN infection. Although a conclusion could not be drawn regarding 16D10 functions based on 16D10 interactions with target proteins, various roles of 16D10 were revealed by the overexpression of this gene in A. thaliana.
The induction of PR5 gene in 16D10-overexpressing lines was repressed in response to
P. syringae. The roots of transgenic lines had more undifferentiated xylem cells and a higher level of VND7 expression. Continuing to study the functional mechanisms of this highly conserved effector in host cells could be useful for understanding the interaction between RKN and host plants, and assist in developing a novel tool to control RKN in crop production.
122
3. 6. ACKNOWLEDGMENTS
The authors thank Drs. Michael Knoblauch, Kiwamu Tanaka and Daniel L.
Mullendore for technical assistance. We also thank to Rachel Olson for technical help.
Funding for this project was provided by grants from the USDA, Washington State
Department of Agriculture, Washington Grain Commission and Northwest Potato
Research Consortium to A. A. Elling. PPNS No. 0672, Department of Plant Pathology,
College of Agricultural, Human, and Natural Resource Sciences, Agricultural Research
Center, Hatch Project No. WNP00744, Washington State University, Pullman, WA
99164.
123
3. 7. LITERATURE CITED
Bendahmane, A., Querci, M., Kanyuka, K., and Baulcombe, D. C. 2000. Agrobacterium
transient expression system as a tool for the isolation of disease resistance
genes: application to the Rx2 locus in potato. Plant J. 21:73-81.
Bird, A. F. 1961. The ultrastructure and histochemistry of a nematode-induced giant cell.
J. Biophys. Biochem. Cytol.11:701-715.
Bird, D. M. K., Opperman, C. H., and Williamson, V. M. 2009. Plant infection by root-
knot nematode. Pages 1-13 in: Cell Biology of Plant Nematode Parasitism. R. H.
Berg, and C. G. Taylor, eds. Berlin and Heidelberg, Springer.
Brigneti, G., Voinnet, O., Li, W. X., Ji, L. H., Ding, S. W., and Baulcombe, D. C. 1998.
Viral pathogenicity determinants are suppressors of transgene silencing in
Nicotiana benthamiana. EMBO J. 17:6739-6746.
Büttner, D., and Bonas, U. 2002. Getting across-bacterial type III effector proteins on
their way to the plant cell. EMBO J. 21:5313-5322.
Caillaud, M. C., Dubreuil, G., Quentin, M., Perfus-Barbeoch, L., Lecomte, P., de
Almeida Engler, J., Abad, P., Rosso, M. N., and Favery, B. 2008. Root-knot
nematodes manipulate plant cell functions during a compatible interaction. J.
Plant Physiol.165:104-113.
Castagnone-Sereno, P. 2002. Genetic variability in parthenogenetic root-knot
nematodes, Meloidogyne spp., and their ability to overcome plant resistance
genes. Nematology 4:605-608.
124
Castagnone-Sereno, P., Bongiovanni, M., and Wajnberg, E. 2007. Selection and
parasite evolution: a reproductive fitness cost associated with virulence in the
parthenogenetic nematode Meloidogyne incognita. Evol. Ecol. 21:259-270.
Chakrabarty, R., Banerjee, R., Chung, S.-M., Farman, M., Citovsky, V., Hogenhout, S.
A., Tzfira, T., and Goodin, M. 2007. pSITE vectors for stable integration or
transient expression of autofluorescent protein fusions in plants: Probing
Nicotiana benthamiana-virus interactions. Mol. Plant-Microbe Interact. 20:740-
750.
Clough, S. J., and Bent, A. F. 1998. Floral dip: a simplified method for Agrobacterium-
mediated transformation of Arabidopsis thaliana. Plant J. 16:735-743.
Cools, T., Iantcheva, A., Weimer, A. K., Boens, S., Takahashi, N., Maes, S., Van den
Daele, H., Van Isterdael, G., Schnittger, A., and De Veylder, L. 2011. The
Arabidopsis thaliana checkpoint kinase WEE1 protects against premature
vascular differentiation during replication stress. Plant Cell 23:1435-1448.
Davies, L. J., Lilley, C. J., Paul Knox, J., and Urwin, P. E. 2012. Syncytia formed by
adult female Heterodera schachtii in Arabidopsis thaliana roots have a distinct
cell wall molecular architecture. New Phytol. 196:238-246.
Davis, E. L., Hussey, R. S., and Baum, T. J. 2004. Getting to the roots of parasitism by
nematodes. TRENDS Parasitol. 20:134-141.
Desai, M., and Hu, J. 2008. Light induces peroxisome proliferation in Arabidopsis
seedlings through the photoreceptor phytochrome A, the transcription factor HY5
HOMOLOG, and the peroxisomal protein PEROXIN11b. Plant Physiol. 146:1117-
1127.
125
Dinh, P. T. Y., Brown, C. R., and Elling, A. A. 2014a. RNA interference of effector gene
Mc16D10L confers resistance against Meloidogyne chitwoodi in Arabidopsis and
potato. Phytopathology 104:1098-1106.
Dinh, P. T. Y., Zhang, L., Brown, C. R., and Elling, A. A. 2014b. Plant-mediated RNA
interference of effector gene Mc16D10L confers resistance against Meloidogyne
chitwoodi in diverse genetic backgrounds of potato and reduces pathogenicity of
nematode offspring. Nematology 16:669-682.
Dinh, P. T. Y., Zhang, L., Mojtahedi, H., Brown, C. R., and Elling, A. A. 2015. Broad
Meloidogyne resistance in potato based on RNA interference of effector gene
16D10. J. Nematol. 47:71-78.
Doyle, E. A., and Lambert, K. N. 2003. Meloidogyne javanica chorismate mutase 1
alters plant cell development. Mol. Plant-Microbe Interact. 16:123-131.
Elling, A. A., Davis, E. L., Hussey, R. S., and Baum, T. J. 2007. Active uptake of cyst
nematode parasitism proteins into the plant cell nucleus. Int. J. Parasitol.
37:1269-1279.
Etchells, J. P., Provost, C. M., Mishra, L., and Turner, S. R. 2013. WOX4 and WOX14
act downstream of the PXY receptor kinase to regulate plant vascular
proliferation independently of any role in vascular organisation. Development
140:2224-2234.
Etchells, J. P., Provost, C. M., and Turner, S. R. 2012. Plant vascular cell division is
maintained by an interaction between PXY and ethylene signalling. PLoS Genet.
8:e1002997.
126
Etchells, J. P., and Turner, S. R. 2010. The PXY-CLE41 receptor ligand pair defines a
multifunctional pathway that controls the rate and orientation of vascular cell
division. Development 137:767-774.
Grebenok, R. J., Pierson, E., Lambert, G. M., Gong, F. C., Afonso, C. L., Haldeman-
Cahill, R., Carrington, J. C., and Galbraith, D. W. 1997. Green-fluorescent protein
fusions for efficient characterization of nuclear targeting. Plant J. 11:573-586.
Godiard, L., Sauviac, L., Torii, K.U., Grenon, O., Mangin, B., Grimsley, N.H. and Marco,
Y. 2003. ERECTA, an LRR receptor-like kinase protein controlling development
pleitropically affects resistance to bacterial wilt. Plant J. 36:353-365.
Haasen, D., Köhler, C., Neuhaus, G., and Merkle, T. 1999. Nuclear export of proteins in
plants: AtXPO1 is the export receptor for leucine-rich nuclear export signals in
Arabidopsis thaliana. Plant J. 20:695-705.
Hassan, S., Behm, C. A., and Mathesius, U. 2010. Effectors of plant parasitic
nematodes that re-program root cell development. Funct. Plant Biol. 37:933-942.
Hewezi, T., Howe, P., Maier, T. R., Hussey, R. S., Mitchum, M. G., Davis, E. L., and
Baum, T. J. 2008. Cellulose binding protein from the parasitic nematode
Heterodera schachtii interacts with Arabidopsis pectin methylesterase:
cooperative cell wall modification during parasitism. Plant Cell 20:3080-3093
Hewezi, T., Howe, P. J., Maier, T. R., Hussey, R. S., Mitchum, M. G., Davis, E. L., and
Baum, T. J. 2010. Arabidopsis spermidine synthase is targeted by an effector
protein of the cyst nematode Heterodera schachtii. Plant Physiol. 152:968-984.
Hewezi, T., Juvale, P. S., Piya, S., Maier, T. R., Rambani, A., Rice, J. H., Mitchum, M.
G., Davis, E. L., Hussey, R. S., and Baum, T. J. 2015. The cyst cematode
127
effector protein 10A07 targets and recruits host posttranslational machinery to
mediate its nuclear trafficking and to promote parasitism in Arabidopsis. Plant
Cell (in press).
Hofmann, J., and Grundler, F. M. W. 2007. Identification of reference genes for qRT-
PCR studies of gene expression in giant cells and syncytia induced in
Arabidopsis thaliana by Meloidogyne incognita and Heterodera schachtii.
Nematology 9:317-323.
Hofmann, J., Youssef-Banora, M., de Almeida-Engler, J., and Grundler, F. M. W. 2010.
The role of callose deposition along plasmodesmata in nematode feeding sites.
Mol. Plant-Microbe Interact. 23:549-557.
Huang, G. Z., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S. 2006a. Engineering
broad root-knot resistance in transgenic plants by RNAi silencing of a conserved
and essential root-knot nematode parasitism gene. Proc. Natl. Acad. Sci. U.S.A.
103:14302-14306.
Huang, G. Z., Dong, R. H., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S.
2006b. A root-knot nematode secretory peptide functions as a ligand for a plant
transcription factor. Mol. Plant-Microbe Interact. 19:463-470.
Hussey, R. S. 1989. Disease-inducing secretions of plant-parasitic nematodes. Annu.
Rev. Phytopathol. 27:123-141.
Hussey, R. S., and Barker, K. R. 1973. A comparison of methods of collecting inocula of
Meloidogyne spp, including a new technique. Plant Dis. Rep. 57:1025-1028.
128
Hussey, R. S., and Mims, C. W. 1990. Ultrastructure of esophageal glands and their
secretory granules in the root-knot nematode Meloidogyne incognita.
Protoplasma 156:9-18.
Iberkleid, I., Vieira, P., de Almeida Engler, J., Firester, K., Spiegel, Y., and Horowitz, S.
B. 2013. Fatty acid-and retinol-binding protein, Mj-FAR-1 induces tomato host
susceptibility to root-knot nematodes. PLoS One 8:e64586.
Jaouannet, M., Magliano, M., Arguel, M. J., Gourgues, M., Evangelisti, E., Abad, P., and
Rosso, M. N. 2013. The root-knot nematode calreticulin Mi-CRT is a key effector
in plant defense suppression. Mol. Plant-Microbe Interact. 26:97-105.
Jaouannet, M., Perfus-Barbeoch, L., Deleury, E., Magliano, M., Engler, G., Vieira, P.,
Danchin, E. G. J., Rocha, M. D., Coquillard, P., Abad, P., and Rosso, M. N. 2012.
A root-knot nematode-secreted protein is injected into giant cells and targeted to
the nuclei. New Phytol. 194:924-931.
Jaouannet, M., and Rosso, M. N. 2013. Effectors of root sedentary nematodes target
diverse plant cell compartments to manipulate plant functions and promote
infection. Plant Signal. Behav. 8:e25507.
Jaubert, S., Milac, A. L., Petrescu, A. J., de Almelda-Engler, J., Abad, P., and Rosso, M.
N. 2005. In planta secretion of a calreticulin by migratory and sedentary stages of
root-knot nematode. Mol. Plant-Microbe Interact. 18:1277-1284.
Jones, M. G. K., and Dropkin, V. H. 1976. Scanning electron microscopy of nematode-
induced giant transfer cells. Cytobios 15:149-161.
129
Kim, M. G., da Cunha, L., McFall, A. J., Belkhadir, Y., DebRoy, S., Dangl, J. L., and
Mackey, D. 2005. Two Pseudomonas syringae type III effectors inhibit RIN4-
regulated basal defense in Arabidopsis. Cell 121:749-759.
Kubo, M., Udagawa, M., Nishikubo, N., Horiguchi, G., Yamaguchi, M., Ito, J., Mimura,
T., Fukuda, H., and Demura, T. 2005. Transcription switches for protoxylem and
metaxylem vessel formation. Genes & Dev. 19:1855-1860.
Lu, S. W., Chen, S. Y., Wang, J. Y., Yu, H., Chronis, D., Mitchum, M. G., and Wang, X.
H. 2009. Structural and functional diversity of CLAVATA3/ESR (CLE)-like genes
from the potato cyst nematode Globodera rostochiensis. Mol. Plant-Microbe
Interact. 22:1128-1142.
Martin, K., Kopperud, K., Chakrabarty, R., Banerjee, R., Brooks, R., and Goodin, M. M.
2009. Transient expression in Nicotiana benthamiana fluorescent marker lines
provides enhanced definition of protein localization, movement and interactions
in planta. Plant J. 59:150-162.
Mitchum, M. G., Wang, X., and Davis, E. L. 2008. Diverse and conserved roles of CLE
peptides. Curr. Opin. Plant Biol. 11:75-81.
Niebel, A., De Almeida Engler, J., Hemerly, A., Ferreira, P., Inzé, D., Van Montagu, M.,
and Gheysen, G. 1996. Induction of cdc2a and cyc1At expression in Arabidopsis
thaliana during early phases of nematode-induced feeding cell formation. Plant J.
10:1037-1043.
Niebel, A., Engler, J. D. A., Tire, C., Engler, G., Van Montagu, M., and Gheysen, G.
1993. Induction patterns of an extensin gene in tobacco upon nematode
infection. Plant Cell 5:1697-1710.
130
Rosso, M.-N., Vieira, P., de Almeida-Engler, J., and Castagnone-Sereno, P. 2011.
Proteins secreted by root-knot nematodes accumulate in the extracellular
compartment during root infection. Plant Signal. Behav. 6:1232-1234.
Sasser, J. N. 1977. Worldwide dissemination and importance of the root-knot
nematodes, Meloidogyne spp. J. Nematol. 9:26-29.
Souza, D. d. S. d. L. e., Junior, J. D. A. d. S., Grossi-de-Sá, M., Rocha, T. L., da Rocha
Fragoso, R., Barbosa, A. E. A. d. D., de Oliveira, G. R., Nakasu, E. Y. T., de
Sousa, B. A., Pires, N. F., Dusi, D. M. d. A., Carneirob, R. M. D. G., Romanob,
E., Almeida-Engler, J. d., Engler, G., Martins-de-Sá, C., and Grossi-de-Sá, M. F.
2011. Ectopic expression of a Meloidogyne incognita dorsal gland protein in
tobacco accelerates the formation of the nematode feeding site. Plant Sci.
180:276-282.
Trudgill, D. L., and Blok, V. C. 2001. Apomictic, polyphagous root-knot nematodes:
exceptionally successful and damaging biotrophic root pathogens. Annu. Rev.
Phytopathol. 39:53-77.
Vieira, P., Danchin, E. G. J., Neveu, C., Crozat, C., Jaubert, S., Hussey, R. S., Engler,
G., Abad, P., de Almeida-Engler, J., Castagnone-Sereno, P., and Rosso, M. N.
2011. The plant apoplasm is an important recipient compartment for nematode
secreted proteins. J. Exp. Bot. 62:1241-1253.
Wang, J., Joshi, S., Korkin, D., and Mitchum, M. G. 2010. Variable domain I of
nematode CLEs directs post-translational targeting of CLE peptides to the
extracellular space. Plant Signal. Behav. 5:1633-1635.
131
Wang, X. H., Mitchum, M. G., Gao, B. L., Li, C. Y., Diab, H., Baum, T. J., Hussey, R. S.,
and Davis, E. L. 2005. A parasitism gene from a plant-parasitic nematode with
function similar to CLAVATA3/ESR (CLE) of Arabidopsis thaliana. Mol. Plant
Pathol. 6:187-191.
Wang, J., Replogle, A., Hussey, R., Baum, T., Wang, X., Davis, E. L., and Mitchum, M.
G. 2011. Identification of potential host plant mimics of CLAVATA3/ESR (CLE)-
like peptides from the plant-parasitic nematode Heterodera schachtii. Mol. Plant
Pathol. 12:177-186.
Whitford, R., Fernandez, A., De Groodt, R., Ortega, E., and Hilson, P. 2008. Plant CLE
peptides from two distinct functional classes synergistically induce division of
vascular cells. Proc. Natl. Acad. Sci. U.S.A. 105:18625-18630.
Williamson, V. M., and Hussey, R. S. 1996. Nematode pathogenesis and resistance in
plants. Plant Cell 8:1735-1745.
Wyss, U., Grundler, F. M. W., and Munch, A. 1992. The parasitic behaviour of second-
stage juveniles of Meloidogyne incognita in roots of Arabidopsis thaliana.
Nematologica 38:98-111.
Xiao, S., and Chye, M.-L. 2011. Overexpression of Arabidopsis ACBP3 enhances
NPR1-dependent plant resistance to Pseudomonas syringe pv tomato DC3000.
Plant Physiol. 156:2069-2081.
Xue, B., Hamamouch, N., Li, C., Huang, G., Hussey, R. S., Baum, T. J., and Davis, E.
L. 2013. The 8D05 parasitism gene of Meloidogyne incognita is required for
successful infection of host roots. Phytopathology 103:175-181.
132
Yamaguchi, M., Kubo, M., Fukuda, H., and Demura, T. 2008. VASCULAR-RELATED
NAC-DOMAIN7 is involved in the differentiation of all types of xylem vessels in
Arabidopsis roots and shoots. Plant J. 55:652-664.
Zhang, L., Davies, L. J., and Elling, A. A. 2014a. A Meloidogyne incognita effector is
imported into the nucleus and exhibits transcriptional activation activity in planta.
Mol. Plant Pathol. 16:48-60.
Zhang, L., Du, L., Shen, C., Yang, Y., and Poovaiah, B. W. 2014b. Regulation of plant
immunity through ubiquitin-mediated modulation of Ca2+-calmodulin-
AtSR1/CAMTA3 signaling. Plant J. 78:269-281.
133
3. 8. FIGURES
Fig. 1. The localization and interaction of
16D10 in Nicotiana benthamiana
Both β-glucuronidase (GUS) and
enhanced green fluorescent protein (eGFP)
were fused at the C/N-terminal of 16D10
mature peptide to create two constructs,
eGFP-GUS-16D10 and 16D10-GUS-eGFP.
These were infiltrated into N. benthamiana to
fluorescently localize 16D10 in the cytoplasm
of tobacco cells as shown in A and B. C to F
show the bimolecular fluorescence
complementation of 16D10 mature peptide with
Arabidosis thaliana target proteins, cytosolic factor (CF, AT1G72150), phosphoinositide specific phospholipase C2 (PLC2 AT3G08510), pfkB-like carbohydrate (PCK,
AT1G19600), and nuclear transport factor 2 (NTF2, AT5G60980). Yellow fluorescent signals from the 16D10-CF and 16D10-PLC2 complexes were localized in the cytoplasm of tobacco cells (C and D respectively). E shows the fluorescent signals from the 16D10-PCK in both cytoplasm and nucleus while F shows the 16D10-NTF2 signals in the intercellular spaces of tobacco cells. Scale bar = 20 µm.
134
Fig. 2. The susceptibility of Arabidopsis
thaliana lines to Meloidogyne incognita
Each plant was inoculated with 250
second-stage juveniles of M. incognita. At 35
days after inoculation (DAI), the M. incognita
egg masses in COL (shown in A) and
OX16D10-1 (shown in B as representative of
the 16D10-overexpressing lines) were stained
with phloxine B. OX16D10-1 had more egg
masses (pink egg masses attached to the
roots) than did COL. After staining at 35 DAI,
the numbers of M. incognita egg masses were counted for each line using six plates/line and three plants/plate (C). The average numbers of egg masses in all 16D10-overexpressing lines, OX16D10-1, OX16D10-2,
OX16D10SP-1 and OX16D10SP-3, were significantly higher than COL (wild type) and
D4 (P < 0.05). D4, 16D10 silencing A. thaliana, had the lowest number of M. incognita egg masses. Each bar represents the mean of egg masses of 18 seedlings per line with standard errors. Letters indicate statistically significant differences using a Student’s t test (P < 0.05). Scale bar = 10 mm.
135
Fig. 3. Pseudomonas syringae DC3000 infection
in Arabidopsis thaliana lines
5 P. syringae DC3000 in 10 mM MgCl2 at 10
concentration were infiltrated into 28 day-old
A. thaliana leaves. At three days after infiltration,
these leaves were weighed and ground in 1 ml of
10 mM MgCl2. The numbers of colony-forming units
(CFU) of P. syringae were determined by serially diluting and plating onto King’s medium B with 50 mg/l of rifampicin and 50 mg/l ampicillin. The average log10 of CFU numbers of P. syringae of all 16D10- overexpressing lines, OX16D10-1, OX16D10-2, OX16D10SP-1 and OX16D10SP-3; the wild type line, COL; and, 16D10 silencing D4 line did not differ significantly (P > 0.05).
Each bar represents the mean of four replicates (seedlings) per line with standard errors.
136
Fig. 4. Callose deposition in
Arabidopsis thaliana lines
One µM flagellin22 (flg22) or sterile
water (control) were infiltrated into 28
day-old A. thaliana leaves. At 24 h after
infiltration, the callose depositions in
these infiltrated leaves were stained with
0.01% aniline blue in phosphate buffer at
pH 9. The callose depositions in COL and
OX16D10-1 leaves in response to flg22,
are shown in A and B respectively. No
callose deposition was found in COL and
OX16D10-1 leaves (shown in C and D,
respectively) infiltrated with sterile water as the control. As shown in E, the average numbers of callose depositions in all 16D10- overexpressing lines, OX16D10-1, OX16D10-2, OX16D10SP-1 and OX16D10SP-3 were significantly higher than in wild type line, COL and 16D10 silencing D4 line (P <
0.05). Each bar represents the mean of six replicates (seedlings) per line with standard errors. Letters indicate statistically significant differences using a Student’s t test (P <
0.05). Scale bar = 200 µm.
137
Fig. 5. The expression level of the PR5
gene encoding pathogen related protein 5
in 16D10-overexpressing Arabidopsis
thaliana lines
P. syringae DC3000 in 10 mM MgCl2 at
105 concentration was infiltrated into 28 day-
old A. thaliana leaves. The RNA samples were
extracted from leaves harvested at 20 h after infiltration. These RNA were used to study the expression of the PR5 gene via quantitative real-time polymerase chain reaction (qRT-PCR). In response to P. syringae, the expression levels of PR5 in all 16D10-overexpressing lines, OX16D10-1, OX16D10-
2, OX16D10SP-1 and OX16D10SP-3, were significantly lower than wild type line, COL and 16D10 silencing D4 line (P < 0.05). Each bar represents the mean of qRT-PCR runs in triplicate with standard errors. Letters indicate statistically significant differences using a Student’s t test (P < 0.05).
138
Fig. 6. The effects of overexpressing
16D10 in Arabidopsis thaliana root
phenotype
The root lengths (in mm) of A. thaliana
lines at 10 days after germination were
measured by ImageJ and are illustrated in A.
The differences in root lengths of 16D10-
overexpressing lines, OX16D10-1, OX16D10-
2, OX16D10SP-1 and OX16D10SP-3; wild
type line, COL; and 16D10 silencing D4 line,
were not consistent. Each bar represents the
mean of 12 replicates (seedlings) per line
with standard errors. Letters indicate
statistically significant differences using a
Student’s t test (P < 0.05). B to G show the
immuno-labelling of the root sections of COL,
OX16D10-1, OX16D10-2, OX16D10SP-1,
OX16D10SP-3 and D4. Transverse sections
of root samples were stained with Calcoflour-
white and immuno-labelled with LM11, a marker of mature xylem cells (white asterisks). OX16D10-1 (C), OX16D10-2 (D),
OX16D10SP-1 (E) and OX16D10SP-3 (F) had more undifferentiated xylem cells (white arrowheads) than COL (B) and D4 (G). Scale bar = 10 µm.
139
Fig. 7. The expression level of the VND7 gene
in 16D10-overexpressing Arabidopsis
thaliana lines
The RNA samples that were extracted
from 10 day-old roots were used to study the
expression of VND7 gene via quantitative real-
time polymerase chain reaction (qRT-PCR). The
expression levels of VND7 in all 16D10- overexpressing lines, OX16D10-1, OX16D10-2, OX16D10SP-1 and OX16D10SP-3, were significantly higher than in wild type line, COL and 16D10 silencing D4 line (P <
0.05). Each bar represents the mean of qRT-PCR runs in triplicate with standard errors.
Letters indicate statistically significant differences using a Student’s t test (P < 0.05).
140
3. 9. SUPPLEMENT Table S1. Primers were used in this study Primer names Primer sequences (5′ – 3′) 16D10 (DQ087264) 16D10-BIF1 TATAGGATCCATGTAAAAATTTAATT 16D10-BIF2 TATAGGATCCATGAAAGCCTAGTGGG 16D10-BIR ATGATGAGCTCTCTTCCTCCAGGATT 16D10-GWF1 GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGGGCAAAAAGCCTAGTG 16D10-GWR1 GGGGACCACTTTGTACAAGAAAGCTGGGTCTCAATTATTTCCTCCAGG 16D10-GWR2 GGGGACCACTTTGTACAAGAAAGCTGGGTCATTATTTCCTCCAGGAT 16D10-EcoRI GGCCGAATTCATGGGCAAAAAGCCTAGTGGGCC 16D10-BamHI-1 CGACGGATCCTCAATTATTTCCTCCAGGATTTGG 16D10-BamHI-2 GCAGGTCGACGGATCCCATTATTTCCTCCAGGATTTGG Cytosolic factor (AT1G72150) CF-GWF GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGGCTCAAGAGGAAGTAC CF-GWR GGGGACCACTTTGTACAAGAAAGCTGGGTCTTGAGTTTTGAACCTGTAGA Nuclear transport factor 2 (AT5G60980) NTF-GWF GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGGCACAGCAGGAAGCTA NTF-GWR GGGGACCACTTTGTACAAGAAAGCTGGGTCTCAAGATGAACCACCACCTC pfkB-like carbohydrate (AT1G19600) PCK-GWF GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGGTAGCTGAGGCCTTG PCK-GWR GGGGACCACTTTGTACAAGAAAGCTGGGTCGTTGTGGATGTCAGGAACA Phosphoinositide-specific phospholipase C2 (AT3G08510) PLC2-GWF GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGTCGAAGCAAACGTAC PLC2-GWR GGGGACCACTTTGTACAAGAAAGCTGGGTCCACAAACTCCACCTTCACG 16D10 (DQ087264) 16D10-RTF GGCAAAAAGCCTAGTGGGCC 16D10-RTR TCAATTATTTCCTCCAGG IRREGULAR XYLEM 1 (IRX1) (AT4G18780) (Cools et al. 2011) IRX1-RTF GCTCGCTGGTCTCGACACAAAT IRX1-RTR GAAGTGACGTCGGAGGGATCAA IRREGULAR XYLEM 3 (IRX3) (AT5G17420) (Cools et al. 2011) IRX3-RTF GGAACGTCGAGCCATGAAGAGA IRX3-RTR GGCCGAGGAAGACTTGGATCAT IRREGULAR XYLEM 5 (IRX5) (AT5G44030) IRX5-RTF CATTCGTCAAAGATCGCAGA IRX5-RTR AGACGTGGCAATTCGTTACC Pathogen-related gene 1 (PR1) (AT2G14160) (Xiao and Chye 2011) PR1-RTF AGGCACGAGGAGCGGTAGG PR1-RTR CATGTTCACGGCGGAGACG Pathogen-related gene 2 (PR2) (AT3G57260) (Xiao and Chye 2011) PR2-RTF TACGGGATGCTAGGCGATACC PR2-RTR CTGGAGGCGAGACGTTCAAGAT Pathogen-related gene 5 (PR5) (AT1G75040) (Xiao and Chye 2011) PR5-RTF TCGGCGATGGAGGATTTGAA PR5-RTR AGCCAGAGTGACGGGAGGAAC VASCULAR-RELATED NAC-DOMAIN 6 (VND6) (AT5G62380) (Kubo et al. 2005) VND6-RTF CCCAACTACAATAATGCAACGA VND6-RTR TTGGCTCATGATTAGCTGAGAA VASCULAR-RELATED NAC-DOMAIN 7 (VND7) (AT1G71930) (Kubo et al. 2005) VND7-RTF GGGACGAATAAAGATCAGAACG VND7-RTR ATGCGGATGTATGACTTGTGTC Ubiquitin carboxyl-terminal hydrolase 22 (AT5G10790) (Hofmann and Grundler 2007 UBP22-RTF ACAACATATGACCCGTTTATCGA UBP22-RTR TGTTTAGGCGGAACGGATACT
141
Fig. S1: Yeast two-hybrid assay (Y2H)
To screen for the host target proteins
that interact with 16D10 effector, Y2H was
performed following the BD Matchmaker
Library Construction and Screening Kits
manual (Clontech Laboratories, Mountain
View, CA). In brief, the mature peptide of
16D10 was inserted into the GAL4-binding domain of pGBKT7 vector. This pGBKT7-16D10 acted as a bait to screen the prey library of A. thaliana cDNA. The library was constructed by cDNA extracted from
A. thaliana roots that were infected by M. incognita. Positive yeast matings, which were screened on selective media (quadruple dropout media, QDO), were validated by co- transformation following manual instructions. Co-transformation yeast cells were dotted onto double dropout media, DDO (as control) and QDO media. All co-transformed yeast colonies were able to grow on DDO. However, only yeast colonies where 16D10 interacted with A. thaliana proteins, nuclear transport factor 2 (NTF2, AT5G60980), cytosolic factor (CF, AT1G72150), phosphoinositide-specific phospholipase C2 (PLC2,
AT3G08510), and pfkB-like carbohydrate (PCK, AT1G19600), were able to grow on the
QDO. SV40 and p53 are control proteins of the Y2H system.
142
Fig. S2: The expression level of 16D10
gene in the 16D10-overexpressing
Arabidopsis thaliana lines
The RNA samples that were extracted
from 10 day-old roots were used to study the
expression of 16D10 gene via quantitative
real-time polymerase chain reaction (qRT-
PCR) with 16D10-RTF and 16D10-RTR primers (Table S1). The expression levels of 16D10 in all 16D10-overexpressing lines,
OX16D10-1, OX16D10-2, OX16D10SP-1 and OX16D10SP-3, as well as with 16D10 silencing D4 line, were significantly higher than the wild type line, COL (P < 0.05). Each bar represents the mean of qRT-PCR runs in triplicate with standard errors. Letters indicate statistically significant differences using a Student’s t test (P < 0.05).
143
CHAPTER FOUR
RNA INTERFERENCE OF EFFECTOR GENE MC16D10L CONFERS RESISTANCE
AGAINST MELOIDOGYNE CHITWOODI IN ARABIDOPSIS AND POTATO
Phuong T.Y. Dinh1, Charles R. Brown2, and Axel A. Elling1
1Department of Plant Pathology, Washington State University, Pullman, WA 99164
2Vegetable and Forage Crops Production Research Unit, United States Department of
Agriculture-Agricultural Research Service, Prosser, WA 99350
Corresponding author : Axel A. Elling
E-mail: [email protected]
This paper was published in 2014 in Phytopathology 104 (10): 1098-1106
GenBank accession number: KF734590
144
4. 1. ABSTRACT
Meloidogyne chitwoodi, a quarantine pathogen, is a significant problem in potato- producing areas worldwide. In spite of considerable genetic diversity in wild potato species, no commercial potato cultivars with resistance to M. chitwoodi are available.
Nematode effector genes are essential for the molecular interactions between root-knot nematodes and their hosts. Stable transgenic lines of Arabidopsis and potato (Solanum tuberosum) with resistance against M. chitwoodi were developed. RNA interference
(RNAi) construct pART27(16D10i-2) was introduced into Arabidopsis thaliana and potato to express double-stranded RNA complementary to the putative M. chitwoodi effector gene Mc16D10L. Plant-mediated RNAi led to a significant level of resistance against M. chitwoodi in Arabidopsis and potato. In transgenic Arabidopsis lines, the number of M. chitwoodi egg masses and eggs was reduced by up to 57 and 67% compared with empty vector controls, respectively. Similarly, in stable transgenic lines of potato, the number of M. chitwoodi egg masses and eggs was reduced by up to 71 and 63% compared with empty vector controls, respectively. The relative transcript level of Mc16D10L was reduced by up to 76% in M. chitwoodi eggs and infective second- stage juveniles that developed on transgenic pART27(16D10i-2) potato, suggesting that the RNAi effect is systemic and heritable in M. chitwoodi.
145
4. 2. INTRODUCTION
The potato (Solanum tuberosum L.) is the most important non-cereal food crop worldwide and makes up the staple diet of over 1 billion people (4,28). Plant-parasitic nematodes are a major threat to potato production worldwide, with root-knot
(Meloidogyne spp.) and cyst nematodes (Globodera spp.) being the most widespread and causing most of the damage (6). In the Pacific Northwest of the United States, which is the leading potato growing area and accounts for more than half of the country’s total production (2), the Columbia root-knot nematode (Meloidogyne chitwoodi
Golden et al.) is the most common and significant nematode threat to sustainable potato cultivation. M. chitwoodi is not only a problem in the United States but has also been found in potato growing regions in Europe, Mexico, Argentina and South Africa, making it a threat in some of the world’s most important potato production areas (21).
M. chitwoodi causes tuber quality defects and can render entire shipments unmarketable (24). It has been estimated that as little as one juvenile per 250 g soil at the beginning of the growing season can lead to a total loss of marketability at harvest
(42). To limit spread, M. chitwoodi has been designated as a quarantine pest, which has a significant impact on the international trade of potato tubers (1).
Meloidogyne spp. are obligate parasites that depend on their host plant for survival. The pathogenic part of the life cycle of root-knot nematodes begins with the second-stage juveniles (J2), which are the infective life stage and invade plant roots, rhizomes and tubers. The J2 migrate intercellularly through host tissue until they become sedentary and induce the formation of giant cells, which constitute the nematodes’ feeding sites and sole source of nutrition. The exact mechanisms that lead
146 to the formation and maintenance of giant cells are unknown, but generally it is thought that secretory proteins from the nematode esophageal gland cells, i.e., effectors, play a key role in the underlying processes (54). After a feeding site has been established, the
J2 increase in size and undergo subsequent molts into third-and fourth-stage juveniles
(J3 and J4). After a final molt they develop into adult females or males. Adult females deposit eggs into egg masses, a gelatinous matrix that protects eggs from desiccation.
Inside the eggs, first-stage juveniles (J1) develop and molt into J2, which hatch under favorable conditions that are mostly dictated by soil moisture and temperature (58).
Nematode control in most potato growing areas is based on routine applications of synthetic nematicides. This practice not only is costly but also potentially harmful to the environment because some products have been linked to negative effects on the atmosphere’s ozone layer (60). Host resistance against root-knot nematodes would be an ideal control strategy, but is difficult to achieve.
In spite of the extremely rich genetic resources found in Solanum sect. Petota, which consists of wild and cultivated potatoes and includes up to 232 tuber-bearing and non-tuber-bearing species (25,27,64,65), exploiting this genetic diversity and introducing M. chitwoodi resistance into cultivated potato (S. tuberosum ssp. tuberosum) has proven to be challenging. Germplasm evaluation identified resistance against M. chitwoodi in a number of potato species, including S. brachistrotrichum
(Bitter) Rydb., S. bulbocastanum Dunal, S. cardiophyllum Lindl., S. chacoense Bitter,
S. fendleri Gray, S. hougasii Corr. and S. stoloniferum Schltdl. (7,9,10,44-46).
Protoplast fusions and sexual hybridization were used to introgress M. chitwoodi resistance from S. bulbocastanum, S. fendleri, S. hougasii and S. stoloniferum into
147 cultivated potato (8,11,12,47) but recent analyses indicate that the same single dominant monogene, RMc1, might be responsible for the resistance found in
S. bulbocastanum, S. fendleri and S. hougasii (15). In spite of over two decades of breeding efforts no commercial potato cultivar with resistance to M. chitwoodi is available to date. The RMc1(blb) gene from S. bulbocastanum has been used to develop advanced S. tuberosum breeding clones, but M. chitwoodi populations are very variable
(34,35) and some isolates are able to overcome the RMc1(blb) gene, presenting a significant problem for traditional resistance breeding (13,48). Furthermore, hairy nightshade (S. sarrachoides Sendtn.), a common weed in potato producing areas in the western United States and a host for M. chitwoodi, was found to undermine resistance under greenhouse and field conditions when it co-occurs with potato breeding lines carrying the RMc1(blb) resistance gene (5).
Given these challenges, transgenic approaches, and in particular the use of RNA interference (RNAi), could be an attractive alternative for nematode control. Plant- parasitic nematodes are known to take up host cytoplasm during the infection process, making them vulnerable to host-derived compounds, including double-stranded RNA
(dsRNA) and small interfering RNA (siRNA). This dependency has been exploited recently by engineering transgenic plant tissue that overexpresses dsRNA that is complementary to nematode genes (for recent reviews see 22,49,51). However, plant- mediated RNAi technology to create nematode resistance has been restricted largely to the model plant Arabidopsis thaliana and Agrobacterium rhizogenes-induced transgenic hairy roots (31,40,63,71). Stable transformation of agriculturally important crops to implement host-mediated RNAi against nematodes has been achieved only in tobacco
148
(Nicotiana tabacum L.) and soybean (Glycine max (L.) Merr.) to date, and with greatly varying levels of resistance (23,66,72). In some of these cases, nematode genes were targeted that are conserved in other animals, and could prove problematic when implemented in the field. Targeting genes that are specific to plant-parasitic nematodes to eliminate off-target effects might provide a better option if this technology is to be employed in crops. The effector gene 16D10 has shown great promise as an RNAi target in other Meloidogyne spp. (31), but it is unknown whether this gene exists in
M. chitwoodi and whether it can be used to create stable resistance against this nematode, especially in agronomically important crops. Therefore, the objective of this study was to test whether creating stable transgenic lines of Arabidopsis and potato that overexpress dsRNA complementary to the Meloidogyne-specific effector gene 16D10 leads to resistance against M. chitwoodi.
4. 3. MATERIALS AND METHODS
Nematode maintenance and extraction. M. chitwoodi isolate WAMC1 was maintained on tomato (S. lycopersicum L.) ‘Rutgers’ under greenhouse conditions (34).
Tomato plants were grown in autoclaved sand. Three months after inoculation,
M. chitwoodi eggs and parasitic life stages were harvested from tomato roots. To extract eggs, roots were cut into small pieces and agitated for 3 min in a 0.5% sodium hypochlorite solution (36). The root suspension was poured over a set of nested test sieves (850, 75, and 25 µm pore size from top to bottom) and eggs were collected from the 25-µm sieve. To purify samples, the M. chitwoodi egg suspension was divided into
50-ml tubes (20 ml per tube) and mixed with 20 ml of 70% sucrose. Ten milliliters of
149 water was carefully placed on top of the suspension and the tubes were immediately centrifuged for 3 min at 375 × g in a clinical centrifuge. After centrifuging, eggs were collected from the interface and rinsed on a 25-µm test sieve. To hatch infective J2, purified eggs were incubated in a modified Baermann pan with sterile water at room temperature for 4 days; hatched infective J2 were collected by centrifugation (19). To extract parasitic nematode life stages, root pieces from which eggs were extracted were blended in tap water and decanted over a set of four nested test sieves (250, 125, 45, and 25 µm pore size from top to bottom). Each sieve was rinsed for 2 min with tap water. M. chitwoodi females were collected from the 125-µm sieve, parasitic J3 and J4 from the 45-µm sieve, and parasitic J2 from the 25-µm sieve, respectively. To purify parasitic nematode stages, samples were transferred to 50-ml tubes and 10 ml of 25%
MgSO4 7H2O was added to the bottom of each tube using a transfer pipet. Tubes were centrifuged as above and parasitic life stages were collected from the interface and rinsed on a 25-µm sieve. To ensure purity of parasitic life stages and to remove remaining plant debris, samples were purified further by manually removing contaminants under a Stemi 2000C stereomicroscope (Zeiss, Jena, Germany). All developmental stages were rinsed either in DEPC-treated water before being stored at
–80°C for use in quantitative real-time polymerase chain reaction (qRT-PCR) or resuspended in 4% formaldehyde in phosphate buffered saline for in situ hybridizations
(see below).
Nematode RNA purification and cloning of Mc16D10L full-length cDNA.
Total RNA was extracted from frozen tissue of all M. chitwoodi life stages using the
PerfectPure RNA Fibrous Tissue kit (5Prime, Gaithersburg, MD) following the
150 manufacturer’s instructions. Approximately 500 ng of purified RNA of each life stage was used as template to synthesize cDNA using the Advantage RT-for-PCR kit
(Clontech, Mountain View, CA) according to the manufacturer’s protocol. Eighty microliters of DEPC-treated water was added to each 20-µl reverse transcription product and samples were stored at –80°C. To clone full-length cDNA of Mc16D10L, primers Mc16D10-F and Mc16D10-R were designed based on previously published
M. chitwoodi expressed sequence tag (EST) sequence CD418743 (61). Using primers
Mc16D10-F and Mc16D10-R and Phusion polymerase (New England Biolabs, Ipswich,
MA), Mc16D10L was amplified from 5 µl of cDNA of infective J2 in a Mastercycler pro thermal cycler (Eppendorf, Hamburg, Germany) with the following PCR program: 94°C for 5 min; 40 cycles at 94°C for 30 s, 52°C for 45 s and 72°C for 30 s, followed by 72°C for 5 min. The PCR product was cloned into pGEM-T Easy (Promega, Madison, WI) and transformed into E. coli DH5-alpha (New England Biolabs). The plasmid was recovered using the GeneJET Plasmid Miniprep kit (Thermo Scientific, Rochester, NY) and sequenced (Elim Biopharm, Hayward, CA) to confirm the identity of the insert.
Sequences were aligned using ClustalW and signal peptides were analyzed with
SignalP 3.0 software.
In situ hybridization. To determine the spatial and temporal expression patterns of Mc16D10L in M. chitwoodi, in situ hybridizations were conducted for all life stages.
After amplifying the Mc16D10L template with primers Mc16D10-F and Mc16D10-R using Phusion polymerase (New England Biolabs), the sense and antisense probes were synthesized with the PCR DIG Probe Synthesis kit (Roche, Indianapolis, IN) following the manufacturer’s instructions. Eggs, infective J2, parasitic J2, J3, J4, and
151 adult females that were fixed previously in 4% formaldehyde in phosphate buffered saline for 2 days at room temperature were cut at random with a razor blade, dehydrated, and hybridized in 1.5-ml microcentrifuge tubes with sense (for negative control) and antisense probes at 37°C for 16 h (37). After incubation, residues were removed and probes were detected following Hussey et al. (37). Hybridization patterns were viewed and documented with an AxioObserver A1 inverted microscope equipped with differential interference contrast, AxioCam ICc1 digital camera, and ZEN imaging software (Zeiss). All experiments were replicated at least twice and showed similar results.
Generation of transgenic Arabidopsis and potato. A. thaliana ecotype
Columbia (Col-0) was used as wild type in infection assays described below and as genetic background for generating transgenic RNAi lines following the floral dip method
(17). Using Agrobacterium tumefaciens strain GV3101, the empty binary vector pART27 and the silencing construct pART27(16D10i-2), which contained a pHANNIBAL RNAi cassette based on Mi16D10 (31), were transformed into A. thaliana. For each construct, approximately 500 seeds from 24 individual plants were germinated on 1/2 strength
Murashige and Skoog (MS) basal salts media (Caisson, North Logan, UT), supplemented with 3% D-sucrose, kanamycin sulfate at 50 µg/ml and timentin at 0.5 g/liter, and solidified with 0.6% Daishin agar at pH 5.7. Seedlings were maintained at
22°C in a growth chamber with a 12 h photoperiod. Selection-resistant seedlings were confirmed by PCR for presence of the transgene, and for each construct, six seedlings were carefully transferred to individual pots filled with potting mix and cultivated in the greenhouse (16 h daylight at 22°C and 8 h nighttime at 20°C). T1 seeds were collected
152 from five individual lines and about 50 T1 seeds for each line were plated on selection media to ensure the presence of the transgene. One plant per line was grown in the greenhouse until maturity to collect T2 seeds, later plated to confirm transgenic phenotypes by kanamycin resistance and PCR. For all experiments described here, T3 lines were used; these were confirmed by PCR and Southern blots (Supplemental
Figure 1). Three transgenic RNAi lines (D1, D2, and D4), one empty vector line (E2), and a wild type control line (Col-0) were used for infection assays.
The same constructs also were introduced into potato ‘Désirée’. Internodal stem pieces were co-incubated with Agrobacterium tumefaciens strain GV3101, carrying the empty pART27 or pART27(16D10i-2) binary vector on sterile CIM media (MS basal salts, 0.25 ppm folic acid, 0.05 ppm D-biotin, 2 ppm glycine, 0.5 ppm nicotinic acid, 0.5 ppm pyridoxine HCl, 0.4 ppm thiamine HCl, 0.01% myo-inositol, 3% D-sucrose, 1 ppm
6-benzylaminopurine, 2 ppm 1-naphthaleneacetic acid, and 0.6% Daishin agar at pH
5.6) for 3 days in the dark at 19°C (14). For the empty pART27 lines, 200 internodal stem pieces were used for transformation, and 250 were used for pART27(16D10i-2), respectively. Stem segments were transferred every 2 weeks to sterile 3C5ZR media
(MS basal salts, 0.5 ppm nicotinic acid, 0.5 ppm pyridoxine HCl, 1 ppm thiamine HCl,
0.01% myo-inositol, 3% D-sucrose, 0.5 ppm indole-3-acetic acid, 3 ppm zeatin ribose, timentin at 0.5 g/liter, kanamycin sulfate at 70 µg/ml, and 0.6% Daishin agar at pH 5.9) and maintained in a growth chamber at 22°C with a 12 h photoperiod (62). Newly formed plantlets that regenerated from callus tissue on 3C5ZR plates were transferred to propagation media (MS basal salts, 3% D-sucrose, kanamycin sulfate at 50 µg/ml, and 0.6% Daishin agar at pH 5.7) and maintained under the same growth conditions (55
153 plantlets for pART27 and 64 plantlets for pART27(16D10i-2)). Regenerated plantlets were analyzed by PCR and northern and Southern blots to ensure presence of the transgene. Three single-insertion pART27(16D10i-2) lines (D54, D56, and D57), one empty vector line (E29), and a wild type control line (DES) were chosen for infection assays in the greenhouse. BLAST searches between the gene-specific region of pART27(16D10i-2) and Arabidopsis and potato ESTs and genome sequences did not yield any significant hits, suggesting that no in planta off-target silencing effects were to be expected.
Infection assays. For Arabidopsis, Col-0 wild type and transgenic (E2, D1, D2, and D4) seeds were surface-sterilized and germinated in a growth chamber (22°C with a 12 h photoperiod) on 1/2 strength MS media supplemented with 3% D-sucrose and kanamycin sulfate at 50 µg/ml (for transgenic lines) and solidified with 0.6% Daishin agar at pH 5.7. After 9 days, seedlings were transferred to individual 500-ml pots filled with SunShine Mix #1 potting mix (Sun Gro Horticulture, Agawam, MA) and grown for
10 days under greenhouse conditions before being inoculated with 5,000 M. chitwoodi eggs per pot. For potato, single nodes of wild type ‘Désirée’ and transgenic (E29, D54,
D56, and D57) lines were cut from plants maintained in tissue culture and propagated in a growth chamber (22°C with a 12 h photoperiod) on MS media supplemented with 3%
D-sucrose and kanamycin sulfate at 50 µg/ml (for transgenic lines) and solidified with
0.6% Daishin agar at pH 5.7 for 1 month. Following the 1-month period, plants were transferred to individual 500-ml pots filled with autoclaved sand, and cultivated under greenhouse conditions for 10 days before being inoculated with 5,000 M. chitwoodi eggs per pot. Sand was used to facilitate cleaning of potato roots at the conclusion of
154 the infection assay. Arabidopsis did not grow well in pure sand, hence the use of potting mix. Both Arabidopsis and potato plants were maintained in the greenhouse for the duration of the experiment, and watered twice per day with 20-10-20 NPK liquid fertilizer. Infection assays were designed as randomized complete block designs with nine replications for each Arabidopsis line and timepoint (35 and 55 days after inoculation [DAI], respectively) and 10 replications for each potato line and timepoint. At
35 DAI, the roots were washed free of growth substrate and for potato, the root fresh weight was measured. Potting mix adhering to roots prevented reliable fresh root weight measurements in Arabidopsis and the weights were omitted. All root systems were stained with phloxine B at 0.15 g/liter for 15 min to visualize M. chitwoodi egg masses as well as facilitate counting under a Stemi 2000C stereomicroscope (Zeiss). At 55 DAI, root systems were washed free of substrate and for potato, root fresh weights were determined as before. For each root system, eggs were extracted as described above and eggs were counted under a stereomicroscope. Eggs were used to extract RNA or hatch infective J2 for qRT-PCR assays as detailed above. All infection assays were conducted twice and showed similar results. Representative results are shown.
Northern blots for transgenic Arabidopsis and potato lines. Total RNA and small RNA were extracted from 0.5 g of leaf tissue of each line using the mirVana miRNA isolation kit (Ambion, Austin, TX). For each line, total RNA (20 µg for
Arabidopsis and 4 µg for potato) was denatured in 50% deionized formamide (Amresco,
Solon, OH), separated on a 1% agarose gel in 1× TBE buffer (0.9 M Tris base, 0.9 M boric acid, and 20 nM EDTA) and transferred by capillary action onto Nytran N nylon membrane (Sigma, St. Louis, MO) in 10× SSC (1.5 M NaCl, 0.15 M sodium citrate, pH
155
7.0). In addition, 1 µg of denatured small RNA was separated on a 15% polyacrylamide gel with 8 M urea in 1× TBE buffer and transferred onto Nytran N membrane as described above. For northern blotting, [α-32P] dATP probes (MP Biochemicals, Solon,
OH) Mi16D10 and U6 were synthesized by PCR using primers 16D10F5, 16D10R5,
U6F, and U6R, respectively (Table 1). Probes were labeled radioactively and purified with DECAprime II kit (Ambion) and Illustra NICK columns (GE Healthcare Life
Sciences, Pittsburgh, PA). Membranes were UV-crosslinked and hybridized with radioactive probes overnight at 37°C in hybridization buffer (50% deionized formamide,
3× SSC, 0.1 mg/ml salmon sperm DNA, 1% sodium dodecyl sulfate, 0.05 M phosphate buffer, 0.2% bovine serum albumin, 0.2% polyvinylpyrrolidone, and 0.2% Ficoll) before being washed three times in 2× SSC and 0.2% SDS for 20 min at 46°C and exposed to
X-ray films for 1 to 5 days at –80°C. All experiments were conducted twice and showed similar results.
qRT-PCR transcript levels of Mc16D10L. To analyze the transcript levels of
Mc16D10L throughout the life cycle of M. chitwoodi, qRT-PCR was performed with cDNA from different nematode life stages harvested from greenhouse nematode stock cultures, as well as from eggs and infective J2 originating from plants used in infection assays (see above). Total RNA was extracted and cDNA generated as described above. qRT-PCR was performed in an iQ Real-Time PCR machine (Bio-Rad, Hercules,
CA) using the iQ SYBR Green Supermix kit (Bio-Rad). Internal transcribed spacer 2
(ITS2) rRNA (JN241865) was chosen as internal control gene for qRT-PCR after comparing the expression pattern with that of other M. chitwoodi housekeeping genes, such as 18S rRNA (AY757835), actin 2 (CB930959), and glyceraldehyde-3-phosphate-
156 dehydrogenase-1 (CB9332930). Primers McITS-RTF and McITS-RTR were designed based on the M. chitwoodi ITS2 gene and used for internal control qRT-PCR reactions
(Table 1). Mc16D10L was amplified using primers Mc16D10-RTF and Mc16D10-RTR
(Table 1). Each sample was run in triplicate using the following program: 94°C for 10 min; 45 cycles at 94°C for 30 s, 50°C for 30 s and 72°C for 30 s, followed by 91 cycles with a temperature increase of 0.5°C after each cycle from 50 to 95°C. The differences in the expression level of Mc16D10L in M. chitwoodi were analyzed using the 2–ΔΔCt method (52) with cycle threshold (Ct) values exported from iQ5 Optical System
Software (Bio-Rad). All experiments were conducted at least twice and showed similar results. Representative results are shown.
Data analysis. Number of egg masses, number of eggs, fold changes, and log10 fold changes of Mc16D10L expression were analyzed in Microsoft Excel to calculate means and standard errors. Statistically significant differences were estimated using a
Student’s t test with alpha = 0.05 in SAS 9.2 (SAS Institute, Cary, NC).
4. 4. RESULTS
M. chitwoodi effector gene Mc16D10L is a ortholog of Mi16D10. In order to identify a putative M. chitwoodi ortholog of the M. incognita effector gene Mi16D10,
M. chitwoodi ESTs were searched using BLASTN (32,61). M. chitwoodi EST CD418743 provided the best match and a full-length sequence spanning the complete open reading frame was cloned from infective J2 cDNA using the gene-specific primers
Mc16D10-F and Mc16D10-R (Table 1). The coding sequence of this Mi16D10-like
M. chitwoodi ortholog, which will be referred to as Mc16D10L henceforth, had a length
157 of 153 bp. A pairwise sequence alignment showed that 70% of the sequences of
Mi16D10 and Mc16D10L were identical on the nucleotide level and 63% on the amino acid level, respectively (Fig. 1). The N-terminal 32 amino acids of Mc16D10L represent a signal peptide, a characteristic of nematode effector gene products, indicating that this putative effector peptide is most likely secreted. Importantly, a region with similarity to the plant CLAVATA3 (CLV3)/ENDOSPERM SURROUNDING REGION (ESR) (CLE) motif (KRXVPXGPNPLHNR) found in Mi16D10 and other Meloidogyne spp. 16D10 orthologs (31,55) also was conserved in Mc16D10L. The nucleotide sequence of
Mc16D10L was deposited in GenBank under accession number KF734590.
Mc16D10L expression changes during nematode development. In situ hybridizations showed that Mc16D10L was expressed most strongly in the subventral gland cells of M. chitwoodi infective J2 (Fig. 2). Weaker expression was found in eggs and subventral gland cells of parasitic J2, whereas no gene activity could be detected in late parasitic life stages by in situ hybridizations. To corroborate the in situ data, qRT-
PCR was performed with cDNA from all M. chitwoodi life stages except adult males
(Fig. 3). Using the expression level of Mc16D10L in eggs as reference, Mc16D10L was upregulated 1.87-fold in infective J2 on a log10 scale, which represents a 78-fold upregulation in absolute terms. In parasitic J2, mixed J3/J4, and adult females,
Mc16D10L was expressed at a lower level than in eggs, with a fold change of –0.72, –1, and –1.6 on a log10 scale (5-fold, 10-fold, and 33-fold downregulation in absolute terms), respectively.
158
Plant-mediated RNAi of Mc16D10L increases M. chitwoodi resistance. No overt phenotypical changes were observed in transgenic Arabidopsis compared with wild type controls. At 35 DAI, the average number of egg masses per plant was reduced
(P < 0.05) by 36 to 45% in RNAi lines D1, D2, and D4 compared with the Col-0 wild type control, and by 50 to 57% compared with the empty vector control, respectively. At 55
DAI, the average number of eggs per plant in Arabidopsis RNAi lines was reduced by
68 to 74% compared with the wild type control, and by 59 to 67% relative to the empty vector control (P < 0.05) (Fig. 4). Since expression of dsRNA of Mc16D10L led to enhanced resistance against M. chitwoodi in the model plant Arabidopsis, the RNAi construct pART27(16D10i-2) was used to generate stable transgenic potato lines to test whether the same strategy could be applied to engineer M. chitwoodi resistance in potato. No overt phenotypical changes were observed in transgenic potato plants compared with wild type controls. At 35 DAI, the average number of egg masses per plant was reduced by 63 to 79% compared with the wild type control, and by 50 to 71% compared with the empty vector control (P < 0.05). Similarly, the average number of eggs per RNAi plant at 55 DAI was reduced by 65 to 74% compared with the wild type, and by 50 to 63% compared with the empty vector control lines (P < 0.05). Comparable results were obtained when nematode infection was measured as number of egg masses or number of eggs per gram of root fresh weight in potato, with reductions of 44 to 56% and 53 to 69% (P < 0.05) compared with the empty vector control, respectively
(Fig. 5). Northern blots confirmed the expression of small RNAs ranging in size from about 50 to over 150 nt in all transgenic pART27(16D10i-2) Arabidopsis and potato lines when a 16D10 probe was used (Fig. 6).
159
Potato RNAi lines downregulate Mc16D10L in M. chitwoodi. To investigate the effect of plant-mediated RNAi on the activity of the target gene in nematodes, qRT-
PCR was used to analyze the relative transcript level of Mc16D10L in M. chitwoodi eggs and J2 harvested from potato plants at 55 DAI. Using the internal ribosomal spacer 2
(ITS2) of nuclear ribosomal DNA as reference (53), it was found that the expression level of Mc16D10L in M. chitwoodi eggs that developed on potato plants carrying pART27(16D10i-2) was reduced by an average of 27 to 76% compared with eggs harvested from wild type control plants (P < 0.05). Similarly, the expression level of
Mc16D10L in infective J2 that hatched from eggs harvested from potato RNAi lines was reduced (P < 0.05) by 52 to 70% relative to J2 that developed from eggs collected from wild type control plants (Fig. 7).
4. 5. DISCUSSION
Plant-mediated downregulation of the putative effector gene Mc16D10L provides resistance against M. chitwoodi in stable transgenic lines of Arabidopsis and potato.
The effector gene 16D10 was identified originally in M. incognita but orthologs have since been found in M. arenaria, M. hapla, and M. javanica (31,33). Whereas 16D10 showed 95 to 98% nucleotide identity and 100% amino acid identity between
M. arenaria, M. hapla, M. incognita, and M. javanica (31), the M. chitwoodi ortholog cloned here was less conserved, with 70% nucleotide and 63% amino acid identity compared with M. incognita. This finding is in line with the more distant phylogenetic position of M. chitwoodi relative to other Meloidogyne spp. in which 16D10 orthologs have been found (30,69). In situ hybridizations and qRT-PCR indicated that Mc16D10L
160 transcription was strongly upregulated in M. chitwoodi infective J2. Even though Huang et al. (33) failed to detect transcripts of Mi16D10 in M. incognita infective J2 by in situ hybridizations, Huang and co-authors later reported that Mi16D10 antiserum confirmed the presence of the Mi16D10 peptide in the subventral glands of M. incognita J2 (32).
This finding suggests that Mi16D10 is in fact actively transcribed in M. incognita infective J2 and possibly escaped detection by Huang and colleagues’ (33) earlier in situ hybridization assays. The function of 16D10 is unknown, but overexpression of the
M. incognita ortholog Mi16D10 increased root growth in tobacco hairy roots and in
A. thaliana (32). All Meloidogyne 16D10 orthologs, including Mc16D10L, contain a region that is similar to the CLE domain of plant CLE signaling peptides, which could mean that this nematode effector has evolved to modulate root growth and vascular development, processes in which CLE peptides play a key role (31,55).
Stable transgenic Arabidopsis and potato lines overexpressing a 271-nt full- length 16D10 dsRNA construct showed strong resistance against M. chitwoodi in this study. No overt changes in plant morphology were observed in the Arabidopsis and potato lines generated in the experiments described here and the level and range of resistance is comparable with what has been reported in other plant-mediated RNAi systems in which cyst or root-knot nematode genes have been targeted (49,51).
Controlling M. chitwoodi under field conditions is challenging due to its wide host range, development at low temperatures and low damage threshold in potato (21). Standard control tactics heavily rely on the application of fumigant and nonfumigant nematicides, but not all compounds and application methods are equally effective at managing
M. chitwoodi (41,42). Alternative control methods, such as green manure tend to reduce
161
M. chitwoodi, but often, the nematode population remains at economically damaging levels or recovers and quickly reaches previous densities once a susceptible crop is planted (56,70). Most likely, nematodes in deeper soil layers are not affected by synthetic nematicides and green manure applied at the soil surface, and enable a rapid infection of a following potato crop (41,70). It is unknown how transgenic 16D10i-2 potatoes would perform under field conditions, but the systemic nature of the resistance suggests that both roots and tubers would be protected at high levels, even in deeper soil layers. Most likely, an ideal M. chitwoodi management system would include diverse approaches, such as traditional nematicides, good crop rotation strategies, green manure, microbial antagonists, and germplasm that carries endogenous or transgenic resistance genes.
Importantly in this study, qRT-PCR indicated a significant reduction in Mc16D10L transcripts in the second-generation M. chitwoodi eggs and infective J2 that developed on the transgenic 16D10i-2 potato lines. A recent report suggests that the RNAi effect of
Mc16D10L in the nematode is systemic and proliferates upon the initial uptake of plant- derived dsRNAs or siRNAs from the esophagus through the entire body of the female, including the gonads and developing eggs, thereby transmitting the RNAi effect to the offspring (20). RNAi is known to be inheritable in Caenorhabditis elegans, making it feasible that the underlying genetic mechanisms are conserved in other nematode taxa
(26). In spite of repeated attempts it was not possible to verify the production of 16D10- specific siRNAs in the Arabidopsis and potato lines created here, a problem that has been encountered previously for other genes, and most likely due to the limited sensitivity of northern blot assays (16,74). Meloidogyne spp. generate a feeding tube
162 during each feeding cycle and it may act as a filter to prevent clogging of the nematode’s mouth spear (38). Earlier studies have shown that root-knot nematodes are able to ingest molecules of 28 to 140 kDa, including the green fluorescent protein and crystal proteins formed by the biocontrol agent Bacillus thuringiensis (50,67,73). It is unknown if M. chitwoodi took up plant-derived dsRNAs and processed them into siRNAs or whether the nematode directly ingested plant-produced siRNAs in this study.
Given that Meloidogyne spp. are able to ingest relatively large molecules, either possibility is conceivable. Previous experiments have demonstrated that a match of 21 nt or less between siRNAs and a target sequence is sufficient to trigger RNAi in animals and that several mismatched base pairs do not interfere with the silencing process
(39,43,57). There are numerous conserved regions between Mi16D10 (upon which pART27(16D10i-2) was designed) and Mc16D10L that fall within this size range and could thereby trigger RNAi in M. chitwoodi.
Almost all commercial potato cultivars are autotetraploid (2n = 4x = 48), which makes classic breeding schemes complicated and time-consuming, especially if the high level of heterozygosity related to tetrasomic inheritance is taken into consideration
(4). Introgressing traits from wild Solanum spp., e.g., nematode resistance genes, compound already challenging breeding strategies and have to overcome additional complications, such as pre- and postzygotic incompatibility barriers (29). Root-knot nematode resistance in wild Solanum spp. is relatively poorly characterized and a largely untapped resource (59). Given the challenges associated with classic potato breeding, transgenic strategies as described here could present an attractive alternative to breed root-knot nematode-resistant potatoes. One of the advantages of creating
163 transgenic plants with an RNAi-based resistance against nematodes is that in principle no foreign protein is expressed in planta, thereby making the end product potentially more desirable than transgenic crops that express nematicidal peptides or proteins, such as cystatins or neurotransmitter antagonists (3,22). Ideal candidate genes for
RNAi-based approaches are specific and only present in the target organisms, even though it has been reported that silencing of host genes can also result in decreased susceptibility against cyst and root-knot nematodes in Arabidopsis (68). Meloidogyne effectors usually lack homology to genes in other taxa, which makes them a worthwhile group of target genes for safe and specific root-knot nematode control (18).
Furthermore, RNAi constructs against more than one effector gene can be stacked to achieve additive effects and aid in creating a more durable nematode resistance (16). In summary, the experiments described here demonstrate that specific silencing of the putative effector gene Mc16D10L leads to M. chitwoodi resistance not only in
Arabidopsis but also in stable transgenic lines of potato, thereby opening the door to improved molecular breeding strategies for nematode resistance in this extremely important food crop.
164
4. 6. ACKNOWLEDGMENTS
We thank R. Hussey and E. Davis for the gift of pART27(16D10i-2) and CSIRO,
Australia for providing the vectors pHANNIBAL and pART27. We thank J. Cardenas for excellent technical assistance. This work was funded by grants from the USDA,
Washington State Department of Agriculture, Washington State Potato Commission,
Idaho Potato Commission, and Washington Grain Commission to A. A. Elling. PPNS
No. 0638, Department of Plant Pathology, College of Agricultural, Human, and Natural
Resource Sciences, Agricultural Research Center, Hatch Project No. WNP00744,
Washington State University, Pullman, WA 99164-6430.
165
4. 7. LITERATURE CITED
1. Anonymous. 2009. Meloidogyne chitwoodi and Meloidogyne fallax. EPPO Bull.
39:5-17.
2. Anonymous. 2011. Potatoes 2010 summary. United States Department of
Agriculture/National Agricultural Statistics Service.
3. Atkinson, H. J., Lilley, C. J., and Urwin, P. E. 2012. Strategies for transgenic
nematode control in developed and developing world crops. Curr. Opin.
Biotechnol. 23:251-256.
4. Barrell, P. J., Meiyalaghan, S., Jacobs, J. M. E., and Conner, A. J. 2013.
Applications of biotechnology and genomics in potato improvement. Plant
Biotechnol. J. 11:907-920.
5. Boydston, R. A., Mojtahedi, H., Brown, C. R., Anderson, T., and Riga, E. 2007.
Hairy nightshade undermines resistance of potato breeding lines to Columbia
root-knot nematode. Am. J. Potato Res. 84:245-251.
6. Brodie, B. B., Evans, K., and Franco, J. 1993. Nematode parasites of potatoes.
Pages 87-132 in: Plant Parasitic Nematodes in Temperate Agriculture. K. Evans,
D. L. Trudgill, and J. M. Webster, eds. CAB International, Wallingford, UK.
7. Brown, C. R., Mojtahedi, H., and Bamberg, J. 2004. Evaluation of Solanum
fendleri as a source of resistance to Meloidogyne chitwoodi. Am. J. Potato Res.
81:415-419.
8. Brown, C. R., Mojtahedi, H., James, S., Novy, R. G., and Love, S. 2006.
Development and evaluation of potato breeding lines with introgressed
166
resistance to Columbia root-knot nematode (Meloidogyne chitwoodi). Am. J.
Potato Res. 83:1-8.
9. Brown, C. R., Mojtahedi, H., and Santo, G. S. 1989. Comparison of reproductive
efficiency of Meloidogyne chitwoodi on Solanum bulbocastanum in soil and in
vitro tests. Plant Dis. 73:957-959.
10. Brown, C. R., Mojtahedi, H., and Santo, G. S. 1991. Resistance to Columbia
root-knot nematode in Solanum spp. and in hybrids of S. hougasii with tetraploid
cultivated potato. Am. Potato J. 68:445-452.
11. Brown, C. R., Mojtahedi, H., and Santo, G. S. 1995. Introgression of resistance to
Columbia and northern root-knot nematodes from Solanum bulbocastanum into
cultivated potato. Euphytica 83:71-78.
12. Brown, C. R., Mojtahedi, H., and Santo, G. S. 1999. Genetic analysis of
resistance to Meloidogyne chitwoodi introgressed from Solanum hougasii into
cultivated potato. J. Nematol. 31:264-271.
13. Brown, C. R., Mojtahedi, H., Zhang, L. H., and Riga, E. 2009. Independent
resistant reactions expressed in root and tuber of potato breeding lines with
introgressed resistance to Meloidogyne chitwoodi. Phytopathology 99:1085-
1089.
14. Brown, C. R., Yang, C. P., Kwiatkowski, S., and Adiwilaga, K. D. 1991. Insert
copy number, chromosome-number, pollen stainability, and crossability of
Agrobacterium-transformed diploid potato. Am. Potato J. 68:317-330.
15. Brown, C. R., Zhang, L., and Mojtahedi, H. 2014. Tracking the RMc1 gene for
resistance to race 1 of Columbia root-knot nematode (Meloidogyne chitwoodi) in
167
three Mexican wild potato species with different ploidies. Am. J. Potato Res.
91:180-185.
16. Charlton, W. L., Harel, H. Y. M., Bakhetia, M., Hibbard, J. K., Atkinson, H. J., and
McPherson, M. J. 2010. Additive effects of plant expressed double-stranded
RNAs on root-knot nematode development. Int. J. Parasitol. 40:855-864.
17. Clough, S. J., and Bent, A. F. 1998. Floral dip: A simplified method for
Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J. 16:735-
743.
18. Danchin, E. G. J., Arguel, M. J., Campan-Fournier, A., Perfus-Barbeoch, L.,
Magliano, M., Rosso, M. N., Da Rocha, M., Da Silva, C., Nottet, N., Labadie, K.,
Guy, J., Artiguenave, F., and Abad, P. 2013. Identification of novel target genes
for safer and more specific control of root-knot nematodes from a pan-genome
mining. PLoS Pathog. 9(10):e1003745.
19. Dinh, P. T. Y., Knoblauch, M., and Elling, A. A. 2014. Nondestructive imaging of
plant-parasitic nematode development and host response to nematode
pathogenesis. Phytopathology 104:497-506.
20. Dinh, P. T. Y., Zhang, L., Brown, C. R., and Elling, A. A. 2014. Plant-mediated
RNA interference of effector gene Mc16D10L confers resistance against
Meloidogyne chitwoodi in diverse genetic backgrounds of potato and reduces
pathogenicity of nematode offspring. Nematology 16:669-682.
21. Elling, A. A. 2013. Major emerging problems with minor Meloidogyne species.
Phytopathology 103:1092-1102.
168
22. Elling, A. A., and Jones, J. T. 2014. Functional characterization of nematode
effectors in plants. Methods Mol. Biol. 1127:113-124.
23. Fairbairn, D. J., Cavallaro, A. S., Bernard, M., Mahalinga-Iyer, J., Graham, M. W.,
and Botella, J. R. 2007. Host-derived RNAi: An effective strategy to silence
genes in plant parasitic nematodes. Planta 226:1525-1533.
24. Finley, A. M. 1981. Histopathology of Meloidogyne chitwoodi (Golden et al.) on
Russet Burbank potato. J. Nematol. 13:486-491.
25. Gavrilenko, T., Antonova, O., Shuvalova, A., Krylova, E., Alpatyeva, N., Spooner,
D. M., and Novikova, L. 2013. Genetic diversity and origin of cultivated potatoes
based on plastid microsatellite polymorphism. Genet. Resour. Crop Evol.
60:1997-2015.
26. Grishok, A., Tabara, H., and Mello, C. C. 2000. Genetic requirements for
inheritance of RNAi in C. elegans. Science 287:2494-2497.
27. Hawkes, J. G. 1990. The Potato: Evolution, Biodiversity and Genetic Resource.
Belhaven Press, London, UK.
28. Hawkes, J. G. 1994. Origins of cultivated potatoes and species relationships.
Pages 3-43 in: Potato Genetics. J. E. Bradshaw and G. R. Mackay, eds. CAB
International, Wallingford, UK.
29. Hermsen, J. G. T. 1994. Introgression of genes from wild species, including
molecular and cellular approaches. Pages 515-538 in: Potato Genetics. J. E.
Bradshaw and G. R. Mackay, eds. CAB International, Wallingford, UK.
30. Holterman, M., Karssen, G., van den Elsen, S., van Megen, H., Bakker, J., and
Helder, H. 2009. Small subunit rDNA-based phylogeny of the Tylenchida sheds
169
light on relationships among some high-impact plant-parasitic nematodes and the
evolution of plant feeding. Phytopathology 99:227-235.
31. Huang, G., Allen, R., Davis, E. L., Baum T. J., and Hussey, R. S. 2006.
Engineering broad root-knot resistance in transgenic plants by RNAi silencing of
a conserved and essential root-knot nematode parasitism gene. Proc. Natl. Acad.
Sci. U.S.A. 103:14302-14306.
32. Huang, G., Dong, R., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S.
2006. A root-knot nematode secretory peptide functions as a ligand for a plant
transcription factor. Mol. Plant-Microbe Interact. 19:463-470.
33. Huang, G., Gao, B., Maier, T., Allen, R., Davis, E. L., Baum, T. J., and Hussey,
R. S. 2003. A profile of putative parasitism genes expressed in the esophageal
gland cells of the root-knot nematode Meloidogyne incognita. Mol. Plant-Microbe
Interact. 16:376-381.
34. Humphreys-Pereira, D. A., and Elling, A. A. 2013. Intraspecific variability and
genetic structure in Meloidogyne chitwoodi from the USA. Nematology 15:315-
327.
35. Humphreys-Pereira, D. A., and Elling, A. A. 2014. Morphological variability in
second-stage juveniles and males of Meloidogyne chitwoodi. Nematology
16:149-162.
36. Hussey, R. S., and Barker, K. R. 1973. A comparison of methods of collecting
inocula of Meloidogyne spp., including a new technique. Plant Dis. Rep. 57:1025-
1028.
170
37. Hussey, R. S., Huang, G., and Allen, R. 2011. Microaspiration of esophageal
gland cells and cDNA library construction for identifying parasitism genes of
plant-parasitic nematodes. Methods Mol. Biol. 712:89-107.
38. Hussey, R. S., and Mims, C. W. 1991. Ultrastructure of feeding tubes formed in
giant-cells induced in plants by the root-knot nematode Meloidogyne incognita.
Protoplasma 162:99-107.
39. Hutvágner, G., McLachlan, J., Pasquinelli, A. E., Bálint, E., Tuschl, T., and
Zamore, P. D. 2001. A cellular function for the RNA-interference enzyme Dicer in
the maturation of the let-7 small temporal RNA. Science 293:834-838.
40. Ibrahim, H. M. M., Alkharouf, N. W., Meyer, S. L. F., Aly, M. A. M., El-Din, A. E.
K. Y. G., Hussein, E. H. A., and Matthews, B. F. 2011. Post-transcriptional gene
silencing of root-knot nematode in transformed soybean roots. Exp. Parasitol.
127:90-99.
41. Ingham, R. E., Hamm, P. B., Baune, M., David, N. L., and Wade, N. M. 2007.
Control of Meloidogyne chitwoodi with shank-injected metam sodium and other
nematicides. J. Nematol. 39:161-168.
42. Ingham, R. E., Hamm, P. B., Williams, R. E., and Swanson, W. H. 2000. Control
of Meloidogyne chitwoodi in potato with fumigant and non-fumigant nematicides.
J. Nematol. (Suppl.) 32:556-565.
43. Jackson, A. L., Bartz, S. R., Schelter, J., Kobayashi, S. V., Burchard, J., Mao, M.,
Li, B., Cavet, G., and Linsley, P. S. 2003. Expression profiling reveals off-target
gene regulation by RNAi. Nat. Biotechnol. 21:635-637.
171
44. Janssen, G. J. W., von Norel, A., Janssen, R., and Hoogendorn, J. 1997.
Dominant and additive resistance to the root-knot nematodes Meloidogyne
chitwoodi and M. fallax in Central American Solanum species. Theor. Appl.
Genet. 94:692-700.
45. Janssen, G. J. W., von Norel, A., Verkerk-Bakker, B., and Janssen, R. 1995.
Detecting resistance to the root-knot nematodes Meloidogyne hapla and M.
chitwoodi in potato and wild Solanum spp. Potato Res. 38:353-362.
46. Janssen, G. J. W., von Norel, A., Verkerk-Bakker, B., and Janssen, R. 1996.
Resistance to Meloidogyne chitwoodi, M. fallax, and M. hapla in wild tuber-
bearing Solanum spp. Euphytica 92:287-294.
47. Janssen, G. J. W., von Norel, A., Verkerk-Bakker, B., Janssen, R., and
Hoogendorn, J. 1997. Introgression of resistance to root-knot nematode from wild
Central American Solanum species into S. tuberosum ssp. tuberosum. Theor.
Appl. Genet. 95:490-496.
48. Janssen, G. J. W., von Norel, A., Verkerk-Bakker, B., Janssen, R., and
Hoogendoorn, J. 1997. Intra- and interspecific variation of root-knot nematodes,
Meloidogyne spp., with regard to resistance in wild tuber-bearing Solanum
species. Fund. Appl. Nematol. 20:449-457.
49. Li, J., Todd, T. C., Lee, J., and Trick, H. N. 2011. Biotechnological application of
functional genomics towards plant-parasitic nematode control. Plant Biotechnol.
J. 9:936-944.
172
50. Li, X. Q., Wei, J. Z., Tan, A., and Aroian, R. V. 2007. Resistance to root-knot
nematode in tomato roots expressing a nematicidal Bacillus thuringiensis crystal
protein. Plant Biotechnol. J. 5:455-464.
51. Lilley, C. J., Davies, L. J., and Urwin, P. E. 2012. RNA interference in plant
parasitic nematodes: A summary of the current status. Parasitology 139:630-640.
52. Livak, K. J., and Schmittgen, T. D. 2001. Analysis of relative gene expression
data using real-time quantitative PCR and the 2–ΔΔCt method. Methods 25:402-
408.
53. McClure, M. A., Nischwitz, C., Skantar, A., Schmitt, M. E., and Subbotin, S. A.
2012. Root-knot nematodes in golf course greens of the western United States.
Plant Dis. 96:635-647.
54. Mitchum, M. G., Hussey, R. S., Baum, T. J., Wang, X., Elling, A. A., Wubben, M.,
and Davis, E. L. 2013. Nematode effector proteins: An emerging paradigm of
parasitism. New Phytol. 199:879-894.
55. Mitchum, M. G., Wang, X., and Davis, E. L. 2008. Diverse and conserved roles of
CLE peptides. Curr. Opin. Plant Biol. 11:75-81.
56. Mojtahedi, H., Santo, G. S., and Ingham, R. E. 1993. Suppression of
Meloidogyne chitwoodi with sudangrass cultivars as green manure. J. Nematol.
25:303-311.
57. Parrish, S., Fleenor, J., Xu, S., Mello, C., and Fire, A. 2000. Functional anatomy
of a dsRNA trigger: Differential requirement for the two trigger strands in RNA
interference. Mol. Cell 6:1077-1087.
173
58. Perry, R. N., Moens, M., and Starr, J. L. (eds.) 2009. Root-Knot Nematodes. CAB
International, Wallingford, UK.
59. Phillips, M. 1994. Inheritance of resistance to nematodes. Pages 319-337 in:
Potato Genetics. J. E. Bradshaw and G. R. Mackay, eds. CAB International,
Wallingford, UK.
60. Porter, I., Banks, J., Mattner, S., and Fraser, P. 2009. Global phaseout of methyl
bromide under the Montreal Protocol: Implications for bioprotection, biosecurity
and the ozone layer. Pages 293-309 in: Recent Developments in Management of
Plant Diseases. Plant Pathology in the 21st Century. U. Gisi, I. Chetand, and M.
L. Gullino, eds. Springer-Verlag, Berlin, Germany.
61. Roze, E., Hanse, B., Mitreva, M., Vanholme, B., Bakker, J., and Smant, G. 2008.
Mining the secretome of the root-knot nematode Meloidogyne chitwoodi for
candidate parasitism genes. Mol. Plant Pathol. 9:1-10.
62. Sheerman, S., and Bevan, M. W. 1988. A rapid transformation method for
Solanum tuberosum using binary Agrobacterium tumefaciens vectors. Plant Cell
Rep. 7:13-16.
63. Sindhu, A. S., Maier, T. R., Mitchum, M. G., Hussey, R. S., Davis, E. L., and
Baum, T. J. 2009. Effective and specific in planta RNAi in cyst nematodes:
Expression interference of four parasitism genes reduces parasitic success. J.
Exp. Bot. 60:315-324.
64. Spooner, D. M. 2009. DNA barcoding will frequently fail in complicated groups:
An example in wild potatoes. Am. J. Bot. 96:1177-1189.
174
65. Spooner, D. M., and Hijmans, R. J. 2001. Potato systematics and germplasm
collecting, 1989–2000. Am. J. Potato Res. 78:237-268.
66. Steeves, R. M., Todd, T. C., Essig, J. S., and Trick, H. N. 2006. Transgenic
soybeans expressing siRNAs specific to a major sperm protein gene suppress
Heterodera glycines reproduction. Funct. Plant Biol. 33:991-999.
67. Urwin, P. E., Møller, S. G., Lilley, C. J., McPherson, M. J., and Atkinson, H. 1997.
Continual green-fluorescent protein monitoring of cauliflower mosaic virus 35S
promoter activity in nematode-induced feeding cells in Arabidopsis thaliana. Mol.
Plant-Microbe Interact. 10:394-400.
68. Van de Cappelle, E., Plovie, E., Kyndt, T., Grunewald, W., Cannoot, B., and
Gheysen, G. 2008. AtCDKA;1 silencing Arabidopsis thaliana reduces
reproduction of sedentary plant-parasitic nematodes. Plant Biotechnol. J. 6:749-
757.
69. van Megen, H., van den Elsen, S., Holterman, M., Karssen, G., Mooyman, P.,
Bongers, T., Holovachov, O., Bakker, J., and Helder, J. 2009. A phylogenetic tree
of nematodes based on about 1200 full-length small subunit ribosomal DNA
sequences. Nematology 11:927-950.
70. Wesemael, W. M. L., and Moens, M. 2008. Vertical distribution of the plant-
parasitic nematode, Meloidogyne chitwoodi, under field crops. Eur. J. Plant
Pathol. 120:249-257.
71. Xue, B., Hamamouch, N., Li, C., Huang, G., Hussey, R. S., Baum, T. J., and
Davis, E. L. 2013. The 8D05 parasitism gene of Meloidogyne incognita is
required for successful infection of host roots. Phytopathology 103:175-181.
175
72. Yadav, B. C., Veluthambi, K., and Subramaniam, K. 2006. Host-generated
double stranded RNA induces RNAi in plant-parasitic nematodes and protects
the host from infection. Mol. Biochem. Parasitol. 148:219-222.
73. Zhang, F., Peng, D., Ye, X., Yu, Z., Hu, Z., Ruan, L., and Sun, M. 2012. In vitro
uptake of 140 kDa Bacillus thuringiensis nematicidal crystal proteins by the
second stage juvenile of Meloidogyne hapla. PLoS ONE 7(6):e38534.
74. Zilberman, D., Cao, X., and Jacobsen, S. E. 2003. ARGONAUTE4 control of
locus-specific siRNA accumulation and DNA histone methylation. Science
299:716-719.
176
4. 8. TABLE
TABLE 1. Primers and probes used for cloning, polymerase chain reaction, and
northern and Southern blots
Gene Primer name Primer sequences (5′ – 3′)
Mi16D10 16D10F5 GTTTACTAATTCAATTAAAAATTTAATT
(DQ087264) 16D10R5 CAATTATTTCCTCCAGGATTTGGCCC
U6 U6F GCGCAAGGATGACACGCA
(X60506) U6R GGCTGAGTTATTTTTTTCTG
ITS2 McITS-RTF GGGGTCAAACCCTTTGGCACGTCTGG
(JN241865) McITS-RTR GCGGGTGATCTCGACTGAGTTCAGG
Mc16D10L Mc16D10-F GATATTTAAATTAAATTATATTCTTCTAAA
(CD418743) Mc16D10-R GCTTTATTCAATTTATTTTTATTTATT
Mc16D10-RTF TTATTTTATCTGTTACTTTTGTGGATTCAGC
Mc16D10-RTR GCGACCATCATTATTATCATTTCCACC
16D10dsRNA 35S-F TTCGCAAGACCCTTCCTCTA
OCS1 CTTCTTCGTCTTACACATCACTTGTC
177
4. 9. FIGURES
Fig. 1. Sequence alignment of 16D10 orthologs from Meloidogyne incognita (Mi16D10) and M. chitwoodi (Mc16D10L). A, Nucleotide level. B, Amino acid level.
Fig. 2. In situ hybridization of Mc16D10L in different Meloidogyne chitwoodi life stages.
Spatial expression pattern of Mc16D10L in A, M. chitwoodi eggs, B, infective second- stage juveniles (J2), and C, parasitic J2. In infective and parasitic J2, Mc16D10L was expressed specifically in the subventral esophageal glands. D, Negative control using a sense probe. Scale bars = 20 µm.
178
Fig. 3. Relative transcript abundance of Mc16D10L in different Meloidogyne chitwoodi life stages. Using the transcript level of Mc16D10L in eggs as a reference, Mc16D10L was upregulated significantly (1.87-fold on a log10 scale) in infective second-stage juveniles (J2). In parasitic J2 (pJ2), mixed J3/J4 parasitic juveniles and adult females
(F), Mc16D10L was downregulated significantly (–0.72, –1.0-, and –1.6-fold on a log10 scale, respectively). Each bar represents the log10 transformed mean of qRT-PCR runs in triplicate with standard errors. Letters indicate statistically significant differences using a Student’s t test (P < 0.05).
179
Fig. 4. Reproductive success of Meloidogyne chitwoodi on transgenic Arabidopsis thaliana expressing pART27(16D10i-2). A, Number of egg masses per plant at 35 days after inoculation (DAI), and B, number of eggs per plant at 55 DAI in Columbia-0 wild type (COL), transgenic empty pART27 vector control (E2), and transgenic pART27(16D10i-2) (D1, D2, and D4) plants. Each bar represents the mean of nine plants per independent line and timepoint with standard errors. Letters indicate statistically significant differences using a Student’s t test (P < 0.05).
180
Fig. 5. Reproductive success of Meloidogyne chitwoodi on transgenic potato expressing pART27(16D10i-2). A, Number of egg masses per plant at 35 days after inoculation
(DAI), B, number of eggs per plant at 55 DAI, C, number of egg masses per gram of root fresh weight at 35 DAI, and D, number of eggs per gram of root fresh weight at 55
DAI in ‘Désirée’ wild type (DES), transgenic empty pART27 vector control (E29), and transgenic pART27(16D10i-2) (D54, D56, and D57) plants. Each bar represents the mean of 10 plants per independent line and timepoint with standard errors. Letters indicate statistically significant differences using a Student’s t test (P < 0.05).
181
Fig. 6. Production of small RNAs in transgenic Arabidopsis and potato plants. A, U6 small nuclear RNA (snRNA) loading control, B, small RNA, and C, total RNA in
Arabidopsis Columbia-0 wild type (COL), transgenic empty pART27 vector control (E2), and transgenic pART27(16D10i-2) (D1, D2, and D4), and in potato ‘Désirée’ wild type
(DES), transgenic empty pART27 vector control (E29), and transgenic pART27(16D10i-
2) (D54, D56, and D57) plants.
182
Fig. 7. Relative fold change of Mc16D10L transcript level in second-generation
Meloidogyne chitwoodi from transgenic potato lines. A, Relative transcript abundance of
Mc16D10L in M. chitwoodi eggs. B, Relative transcript abundance of Mc16D10L in
M. chitwoodi infective second-stage juveniles (J2). Eggs were harvested from transgenic potato lines expressing the silencing construct pART27(16D10i-2) and used either directly for qRT-PCR or allowed to hatch infective J2 in modified Baermann pans.
Potato lines included ‘Désirée’ wild type (DES), transgenic empty pART27 vector control
(E29), and transgenic pART27(16D10i-2) (D54, D56, and D57). Each bar represents the mean of qRT-PCR runs in triplicate with standard errors. Letters indicate statistically significant differences using a Student’s t test (P < 0.05).
183
Supplemental Fig. 1. Southern blot for transgenic Arabidopsis and potato. Lines used include Arabidopsis Columbia-0 wild type (COL), transgenic empty pART27 vector control (E2) and transgenic pART27(16D10i-2) (D1, D2, D4), as well as potato ‘Désirée’ wild type (DES), transgenic empty pART27 vector control (E29) and transgenic pART27(16D10i-2) (D54, D56, D57). Fifteen µg genomic DNA of each Arabidopsis and potato line was digested with XbaI (New England Biolabs), loaded on a 0.8% agarose gel, transferred to a membrane and hybridized with a [α-32P] dATP (MP Biochemicals) probe that was synthesized with primers 35S-F and OCS1 (Table 1). None of the wild type or empty vector control lines showed any signal. Lines D1 and D2 had multiple insertions of the transgene, whereas lines D4, D54, D56 and D57 had single insertions.
184
CHAPTER FIVE
PLANT-MEDIATED RNA INTERFERENCE OF EFFECTOR GENE MC16D10L
CONFERS RESISTANCE AGAINST MELOIDOGYNE CHITWOODI IN
DIVERSE GENETIC BACKGROUNDS OF POTATO AND REDUCES
PATHOGENICITY OF NEMATODE OFFSPRING
Phuong T.Y. Dinh1, Linhai Zhang2, Charles R. Brown2, and Axel A. Elling1
1Department of Plant Pathology, Washington State University, Pullman, WA 99164
2United States Department of Agriculture-Agricultural Research Service, Prosser, WA
99350
Corresponding author : Axel A. Elling
E-mail: [email protected]
This paper was published in 2014 in Nematology 16 (6): 669-682
Keywords: Columbia root-knot nematode, molecular breeding, RNAi, transgenic, transmission, Solanum tuberosum
185
5. 1. ABSTRACT
Meloidogyne chitwoodi is a major problem for potato production in the Pacific
Northwest of the USA. In spite of long-term breeding efforts no commercial potato cultivars with resistance to M. chitwoodi exist to date. The RMc1(blb) resistance gene against M. chitwoodi has been introgressed from Solanum bulbocastanum into cultivated potato (S. tuberosum), but M. chitwoodi pathotypes are able to overcome this resistance. In this study, an RNA interference (RNAi) transgene targeting the
M. chitwoodi effector gene Mc16D10L was introduced into potato cvs Russet Burbank and Désirée, and the advanced breeding line PA99N82-4, which carries the RMc1(blb) gene. Stable transgenic lines were generated for glasshouse infection assays. At 35 days after inoculation (DAI) with M. chitwoodi race 1 the number of egg masses (g root)−1 formed on RNAi lines of cvs Russet Burbank and Désirée was reduced significantly by up to 68% compared to empty vector control plants. At 55 DAI, the number of eggs was reduced significantly by up to 65%. In addition, RNAi of Mc16D10L significantly reduced the development of egg masses and eggs formed by the RMc1(blb) resistance-breaking M. chitwoodi pathotype Roza on PA99N82-4 by up to 47 and 44%, respectively. Importantly, the plant-mediated silencing effect of Mc16D10L was transmitted to M. chitwoodi offspring and significantly reduced pathogenicity in the absence of selection pressure on empty vector control plants. This finding suggests that the RNAi effect is stable and nematode infection decreases regardless of the genotype of the host once the RNAi process has been initiated in the nematode through a transgenic plant. In summary, plant-mediated down-regulation of effector gene
Mc16D10L provides a promising new tool for molecular breeding against M. chitwoodi.
186
5. 2. INTRODUCTION
Root-knot nematodes (Meloidogyne spp.) are among the most damaging plant pathogens worldwide, and infect almost all vascular plants. They are obligate endoparasites that depend on their hosts for survival. Second-stage juveniles (J2), which are the infective life stage, migrate intercellularly through root tissue until they become sedentary and establish a feeding site. The exact molecular mechanisms that define plant-nematode interactions and ultimately lead to the formation and maintenance of feeding sites are unknown, but it seems almost certain that proteinaceous secretions (i.e., effectors) from the pharyngeal gland cells of the nematode play an essential role in the process (Mitchum et al., 2013). After a feeding site has been established, the J2 develop into third- and fourth-stage juveniles, separated by moults. After a fourth and final moult, the nematodes turn into adult females or males. The adult males regain their motility, leave the root and fertilise the females. By contrast, adult females remain sedentary and deposit eggs into egg masses, gelatinous matrices that protect eggs from desiccation. Inside the eggs, first- stage juveniles develop and moult into J2, which hatch under favourable conditions (see
Perry et al., 2009).
Potatoes (Solanum tuberosum) are the most important non-cereal food crop and represent the staple diet for over 1 billion people (Hawkes, 1994; Barrell et al., 2013).
Total annual production on a global scale has been estimated at over 374 million metric tons, with the USA and western Europe producing the highest yields of about 50 t ha−1
(Anon., 2012; Barrell et al., 2013). Plant-parasitic nematodes, especially Meloidogyne spp. and Globodera spp., are a major problem in most potato-growing regions (Brodie
187 et al., 1993). In the Pacific Northwest, which accounts for more than half of the total potato production in the USA, the Columbia root-knot nematode (M. chitwoodi) is of particular concern (Anon., 2011). Meloidogyne chitwoodi is widespread and causes tuber quality defects that can render entire shipments unmarketable (Finley, 1981;
Elling, 2013). The economic damage threshold of this nematode species is exceptionally low. It has been estimated that as few as 1 J2 (250 g soil)−1 at the beginning of the growing season can cause a total loss of marketability at harvest
(Ingham et al., 2000). Meloidogyne chitwoodi has been declared a quarantine pest and complicates the international trade of infested tubers (Anon., 2009).
Nematode control in most potato-growing regions depends heavily on the application of synthetic nematicides, but this practice can be costly and harmful to the environment (Porter et al., 2009). Host resistance would be an ideal control strategy, and testing of germplasm has identified M. chitwoodi resistance in the wild potato species, S. brachistrotrichum, S. bulbocastanum, S. cardiophyllum, S. chacoense,
S. fendleri, S. hougasii and S. stoloniferum (Brown et al., 1989, 1991a, 2004; Janssen et al., 1995, 1996, 1997). A single dominant monogene, RMc1, might be responsible for the resistant phenotype in S. bulbocastanum, S. fendleri and S. hougasii (Brown et al.,
2013). However, in spite of the extremely rich genetic resources found in wild potato species, no commercial potato cultivars with resistance to M. chitwoodi exist to date.
Solanum bulbocastanum has been used to develop introgression lines of cultivated potatoes that are resistant to M. chitwoodi, but specific M. chitwoodi pathotypes that are able to overcome RMc1(blb)-mediated resistance are present in the Pacific Northwest
(Brown et al., 1995, 2006, 2009). Pathotypes are defined as populations that reproduce
188 on a host species that is resistant to other populations of the same nematode species, whereas races are defined as intraspecific variants that can be separated by their ability to reproduce on different plant genera (Dropkin, 1988). Thus, given these pathotypes it seems unlikely that RMc1(blb)-carrying advanced breeding lines would be effective at fully controlling M. chitwoodi.
Most commercial potato cultivars are autotetraploid, which makes potato breeding and genetics exceptionally complicated. Biotechnological advances could provide a feasible option to overcome some of these challenges. RNA interference
(RNAi)-induced silencing offers a promising new technology for engineering a variety of traits in plants and animals. RNAi was first described in Caenorhabditis elegans and now is recognised as a universal phenomenon in a wide range of organisms (Fire et al.,
1998). The basic mechanism involves the processing of large double-stranded RNA
(dsRNA) molecules into shorter double-stranded fragments of small interfering RNA
(siRNA) that are about 21 nt long. Processing is catalysed by Dicer, a ribosome III-like enzyme. Each siRNA is unwound into two separate strands, named passenger and guide. The passenger strand is degraded, and the guide strand is loaded into RISC
(RNA induced silencing complex), a multisubunit complex. Gene silencing occurs when the guide strand binds to a complementary region of target gene mRNA, which is in turn degraded (Hammond et al., 2001). During plant infection, Meloidogyne spp. take up cytoplasm from host plant cells and are capable of ingesting molecules of 28-140 kDa
(Urwin et al., 1997; Li et al., 2007; Zhang et al., 2012). This interaction provides an intriguing possibility to create nematode resistance by engineering transgenic plants that produce dsRNAs and/or siRNAs complementary to nematode genes. Upon uptake
189 of small RNAs through the plant cytoplasm, the endogenous RNAi machinery of the nematode leads to degradation of the complementary mRNA and results in down- regulation or deactivation of the target gene (Li et al., 2011). Effector genes play an essential role for plant-nematode interactions. This makes them an attractive group of genes for RNAi-induced down-regulation, not only because they are crucial for nematode survival, but also because they tend to lack significant homology to genes in other organisms, thereby avoiding off-target effects.
The Meloidogyne-specific effector gene 16D10 has proven to be an especially effective target to control root-knot nematodes. The function of 16D10 is unknown but it encodes a region that has similarity to plant CLAVATA3 (CLV3)/endosperm surrounding region (ESR) (CLE) motifs, i.e., peptides that are involved in controlling root architecture
(Huang et al., 2006a; Mitchum et al., 2008). Overexpression of 16D10 led to increased root growth in tobacco (Nicotiana tabacum) hairy roots and Arabidopsis thaliana (Huang et al., 2006a). In another study, an RNAi construct overexpressing a 271 nt full-length dsRNA region of 16D10 was introduced into A. thaliana. Inoculations with M. arenaria,
M. hapla, M. incognita and M. javanica resulted in a reduction of the number of galls and eggs by 63-90% and 69-93%, respectively (Huang et al., 2006b). Further, a 16D10-
RNAi construct was introduced into hairy roots of grape (Vitis vinifera) to engineer M. incognita resistance. However, the results of that study were inconclusive because hairy root morphology might have been a strong confounding factor, with the thickest roots showing the least nematode infection (Yang et al., 2013). Most recently, Mc16D10L, a
16D10 orthologue was identified in M. chitwoodi, and a 16D10-RNAi construct shown to
190 lead to dramatically increased levels of M. chitwoodi resistance in A. thaliana and potato
(Dinh et al., 2014).
In this report, RNAi technology was used to develop M. chitwoodi resistance in potato cvs Russet Burbank and Désirée and advanced breeding line PA99N82-4.
Russet Burbank makes up about 40% of the total potato acreage in the USA, and
Désirée is an important specialty cultivar, particularly in Europe (Anon., 2013).
PA99N82-4 carries the RMc1(blb) resistance gene from S. bulbocastanum (Brown et al.,
2009). The objectives of this study were: i) to analyse whether the genetic background of potato has an effect on 16D10-RNAi-mediated M. chitwoodi resistance; ii) to test whether 16D10-RNAi resistance is effective against the RMc1(blb)-breaking M. chitwoodi pathotype Roza; and iii) to analyse if the effect of the 16D10-RNAi reduces the pathogenicity of M. chitwoodi offspring to potato.
5. 3. MATERIALS AND METHODS
GENERATION OF TRANSGENIC POTATO LINES
Silencing construct pART27(16D10i-2) was designed previously to down-regulate the Mc16D10L ortholog Mi16D10 in M. incognita (Huang et al., 2006b). The coding sequences of Mi16D10 (AY134435) and Mc16D10L (KF734590) are 70% identical on the nucleotide level and 63% on the amino-acid level, respectively. Internodal stem segments of potato cvs Désirée and Russet Burbank, as well as advanced breeding line
PA99N82-4, were co-incubated with Agrobacterium tumefaciens strain GV3101 carrying the RNAi silencing construct pART27(16D10i-2) or empty vector control pART27
(Brown et al., 2006; Huang et al., 2006b). After 3 days on CIM media (MS basal salts,
191
0.25 ppm folic acid, 0.05 ppm D-biotin, 2 ppm glycine, 0.5 ppm nicotinic acid, 0.5 ppm pyridoxine HCl, 0.4 ppm thiamine HCl, 0.01% myo-inositol, 3% D-sucrose, 1 ppm 6- benzylaminopurine, 2 ppm 1-naphthaleneacetic acid, 0.6% Daishin agar at pH 5.6), stem segments were transferred to 3C5ZR media (MS basal salts, 0.5 ppm nicotinic acid, 0.5 ppm pyridoxine HCl, 1 ppm thiamine HCl, 0.01% myo-inositol, 3% D-sucrose,
0.5 ppm indole-3-acetic acid, 3 ppm zeatin ribose, 0.5 g l−1 timentin, 70 µg ml−1 kanamycin sulphate, 0.6% Daishin agar at pH 5.9) and incubated in a growth chamber
(22°C, 12 h photoperiod) for about 3 months until shoots developed (Sheerman &
Bevan, 1988; Brown et al., 1991b). Stem segments were transferred to fresh 3C5ZR media every 2 weeks. Potato plantlets that regenerated on 3C5ZR were maintained on propagation media (MS basal salts, supplemented with 3% D-sucrose, 50 µg ml−1 kanamycin sulphate and 50 µg ml−1 timentin, solidified with 0.6% Daishin agar) in a growth chamber at 22°C with a 12 h photoperiod. Wild type lines of cvs Désirée and
Russet Burbank and PA99N82-4 also were treated the same way, except that the propagation media did not contain kanamycin sulphate and timentin.
DNA EXTRACTION AND SOUTHERN BLOTTING OF TRANSGENIC POTATO LINES
DNA was extracted from leaves of all putative pART27(16D10i-2) and pART27 transformants that survived kanamycin selection. DNA extracted from wild type cvs
Désirée and Russet Burbank and advanced breeding line PA99N82-4 served as controls. Approximately 1 g leaf material was ground in liquid nitrogen and homogenized in 3 ml TPS extraction buffer (100 mM Tris-HCl (pH 8.0), 100 mM EDTA
(pH 8.0), 1 M KCl). The resulting leaf extraction suspensions were transferred to 15 ml tubes, incubated at 75°C for 10 min, and centrifuged at 13 500 g for 10 min.
192
Supernatants were collected and mixed with an equal volume of isopropanol by inverting each tube several times. DNA was pelleted by centrifuging at 2500 g for 10 min, washed with 70% ethanol, air dried, resuspended in 0.4 ml sterile water containing
0.5 mg ml−1 RNase (Fermentas) and incubated at 37°C for 30 min. After RNase treatment, DNA samples were mixed with 0.4 ml chloroform, vigorously shaken for 1 min and centrifuged at 13 500 g for 5 min. After centrifugation, the top layer of each tube was collected and mixed with an equal volume of isopropanol by inverting each tube several times. Samples were centrifuged at 18 000 g for 10 min, and pelleted DNA was washed with 70% ethanol, air dried and resuspended in 100 µl sterile water.
Prior to Southern blotting, DNA from putative transformants was analysed by
PCR for presence of the transgene, using primers 35S-F and OCS1 for pART27(16D10i-2) and primers nptII-F and nptII-R for pART27, respectively (data not shown). For Southern blots, 15 µg DNA of each PCR-positive potato line was digested with 50 U XbaI (New England Biolabs) for 16 h at 37°C. Digested DNA was separated on a 0.8% agarose gel at 70 V for 16 h before being transferred by capillary action to a
GeneScreen Plus nylon membrane (PerkinElmer) in 10× saline sodium citrate (SSC) buffer (1.5 M NaCl, 0.15 M sodium citrate, pH 7.0). The membranes were cross-linked by UV, and hybridised overnight in hybridization buffer (50% deionised formamide, 0.1 mg ml−1 salmon sperm DNA, 1% sodium dodecyl sulphate (SDS), 1 M NaCl, 10% dextran sulphate) at 42°C with probe 16D10i-2. The probe was amplified with primers
35S-F and OCS1 using plasmid pART27(16D10i-2) as template, radioactively labelled with [α-32P]dATP (MP Biochemicals) using the DECAprime II kit (Ambion) and purified with Illustra NICK columns (GE Healthcare Life Sciences). After hybridisation, the
193 membranes were washed twice with 2× SSC buffer for 5 min at 42°C, followed by three washes with 2× SSC plus 1% SDS for 20 min at 65°C. Three final washes were carried out with 0.1× SSC plus 1% SDS for 20 min each at 42°C, after which the membranes were exposed to X-ray films (Research Products International) for 2 days at −80°C. All experiments were conducted twice, and showed similar results.
NORTHERN BLOTTING OF TRANSGENIC POTATO LINES
Total RNA enriched with small RNAs was extracted from 1 g of leaves using the mirVana miRNA isolation kit (Ambion). For each potato line, 20 µg denatured total RNA was separated on a 1% agarose gel at 120 V for 1 h before being transferred by capillary action to Nytran N nylon membranes (Sigma-Aldrich) overnight. Probes 16D10 and U6 were synthesised using primers 16D10F5, 16D10R5, U6F and U6R (Table 1), respectively, with plasmid pART27(16D10i-2) and potato cDNA serving as templates.
Probes were radioactively labelled with [α-32P]dATP (MP Biochemicals) as described above, and used to hybridise the membranes overnight at 25°C in hybridisation buffer
(50% deionised formamide, 3× SSC, 0.1 mg ml−1 salmon sperm DNA, 1% SDS, 0.05 M phosphate buffer, 0.2% bovine serum albumin, 0.2% polyvinylpyrrolidone, 0.2% Ficoll).
After hybridisation, membranes were washed three times with 2× SSC plus 0.2% SDS for 20 min at 46°C before being exposed to X-ray films (Research Products
International) for 1-7 days at −80°C. All experiments were conducted at least three times and showed similar results.
NEMATODE INOCULUM AND INFECTION ASSAYS
Meloidogyne chitwoodi isolates WAMC1 (race 1) and Roza (race 1, pathotype
Roza) were maintained on tomato (S. lycopersicum) cv. Rutgers under glasshouse
194 conditions. To ensure isolate purity, a portion of each batch of inoculum was used in an assay with indicator host plants as described previously (Brown et al., 2009;
Humphreys-Pereira & Elling, 2013). To obtain nematode inoculum, M. chitwoodi eggs were collected from tomato plants that were inoculated about 3 months earlier following routine procedures (Hussey & Barker, 1973). Briefly, infected roots were cut into 2-3 cm pieces and shaken in 0.5% NaOCl for 3 min. The root suspension was poured over a set of nested test sieves (850, 75, 25 µm pore size from top to bottom) and eggs were collected on the 25 µm pore sieve. For nematode offspring RNAi infection assays (see below), M. chitwoodi WAMC1 eggs were collected from potato cv. Désirée lines E29
(carrying empty vector pART27) and D56 (carrying RNAi silencing construct pART27(16D10i-2)) that were inoculated with M. chitwoodi eggs 4 months earlier and maintained under glasshouse conditions. Egg extractions were the same as described above.
Transgenic pART27(16D10i-2) potato lines, with cv. Désirée (D56, D57, D12 and
D42), cv. Russet Burbank (D5, D16, D20 and D25) and PA99N82-4 (D2, D17, D53 and
D55) as genetic backgrounds, were chosen for infection assays based on transgene copy numbers and overall phenotypic appearance. Single nodes were cut from each line and maintained for 1 month on propagation media supplemented with 50 µg ml−1 kanamycin sulphate. Similarly, single nodes of wild type and pART27 empty vector control lines of each genetic background (DES and E29 for cv. Désirée, RB and E34 for cv. Russet Burbank, 82-4 and E12 for PA99N82-4, respectively) were cut and maintained on propagation media lacking kanamycin sulphate for an equal amount of time. One month-old plantlets were transferred to individual Ray Leach SC10U cone-
195 tainers (Stuewe & Sons) filled with autoclaved sand. Cone-tainers were placed in RL98 trays (Stuewe & Sons) and plants were allowed to acclimate to glasshouse conditions for 10 days, after which each cone-tainer was inoculated with 2000 M. chitwoodi eggs.
Plants were maintained in the glasshouse for the duration of the experiment, and watered twice a day with 20-10-20 NPK liquid fertiliser. Infection assays were set up as randomized complete block designs with 10 replicates (10 plants) per line and time- point (35 and 55 days after inoculation, DAI). All experiments were conducted twice and showed similar results. Three distinct plant/nematode combinations were analysed: i) potato cv. Désirée, cv. Russet Burbank and PA99N82-4 inoculated with M. chitwoodi isolate WAMC1; ii) PA99N82-4 inoculated with M. chitwoodi isolate Roza; and iii) cv.
Désirée inoculated with M. chitwoodi isolate WAMC1 collected from potato lines with and without the 16D10i-2 transgene. For i) and ii), nematode inoculum was collected from tomato cv. Rutgers as described above and the infection assays were conducted in the glasshouse. For iii), which was designed to test the RNAi effect on nematode offspring, M. chitwoodi inoculum was harvested from transgenic potato cv. Désirée lines
E29 (empty vector control) and D56 (carrying pART27(16D10i-2)), resulting in four different treatments: empty vector line E29 inoculated with M. chitwoodi eggs collected from line E29 (pE29-eE29); empty vector line E29 inoculated with M. chitwoodi eggs collected from 16D10i-2 line D56 (pE29-eD56); 16D10i-2 line D56 inoculated with
M. chitwoodi eggs collected from empty vector line E29 (pD56-eE29); and 16D10i-2 line
D56 inoculated with M. chitwoodi eggs collected from line D56 (pD56-eD56), where ‘p’ stands for plant and ‘e’ stands for eggs. pE29 and pD56 served as additional controls, and were inoculated with M. chitwoodi WAMC1 eggs collected from wild type tomatoes
196 cv. Rutgers. Eggs harvested from pE29 and pD56 served as inoculum for the infection assays in iii). Nematode infection assays for iii) were conducted in a growth chamber
(25°C day, 21°C night, 16 h photoperiod).
For each plant/nematode combination and experiment, ten plants were harvested at 35 DAI, their roots were washed free of sand and the fresh weight of roots was determined. Root systems were stained with 0.15 g l−1 phloxine B for 15 min to visualise egg masses, and facilitate counting under a stereomicroscope. At 55 DAI, an additional ten plants were harvested, the roots washed, and root fresh weight determined as before. Eggs were extracted as described above, and again counted under a stereomicroscope.
NEMATODE RNA EXTRACTION, QUANTITATIVE REAL-TIME PCR AND NORTHERN BLOTS
For infection assay iii) (see above) an aliquot of each batch of eggs harvested from experimental plants at 55 DAI was used to hatch J2 in a modified Baermann pan.
J2 were collected by centrifugation and flash frozen in liquid nitrogen. Total RNA was extracted from J2 with the PerfectPure RNA Fibrous Tissue kit (5Prime). For qRT-PCR,
500 ng total RNA was used to synthesise a total volume of 100 µl cDNA with the
Advantage RT-for-PCR kit (Clontech). qRT-PCR for analysing the transcript level of
Mc16D10L was conducted in an iQ Real-Time PCR machine with iQ SYBR Green
Supermix (Bio-Rad). Primers Mc16D10-RTF and Mc16D10-RTR were used to amplify target gene Mc16D10L. Internal transcribed spacer 2 (ITS2) rRNA (JN241865) served as the control, and was amplified with primers McITS-RTF and McITS-RTR (Table 1).
Differences in transcript levels were analysed using the 2–ΔΔCt method (Livak &
197
Schmittgen, 2001) with Ct values retrieved from iQ5 Optical System Software (Bio-Rad).
All reactions were run in triplicate, conducted twice, and gave similar results.
For Northern blots, RNA was transferred to Nytran N membranes (Sigma-Aldrich) and hybridised with probe Mc16D10L (Table 1), generated with primers Mc16D10-F and
Mc16D10-R, and radioactively labelled with [α-32P]dATP (MP Biochemicals) as described above. As the control, a probe for ITS2 was generated with primers McITS-
RTF and McITS-RTR (Table 1).
DATA ANALYSIS
Number of egg masses, number of eggs and relative fold changes of Mc16D10L transcript levels were analysed in Microsoft Excel to calculate means and standard errors. Statistically significant differences were estimated in SAS 9.2 using a Student’s t
-test with P = 0.05.
5. 4. RESULTS
RNAI TRANSGENE 16D10I-2 INCREASES RESISTANCE AGAINST M. CHITWOODI IN DIFFERENT
GENETIC BACKGROUNDS OF POTATO
Stable transgenic lines of cv. Désirée, cv. Russet Burbank and PA99N82-4 each carrying the silencing construct pART27(16D10i-2) were generated to test whether the genetic background of potato has an effect on plant-mediated RNAi resistance against
M. chitwoodi. Empty vector lines transformed with pART27 and wild type plants served as controls. No overt phenotypical changes were observed in transgenic plants compared to wild type controls throughout development. To examine whether the copy number of the RNAi transgene is related to the level of nematode resistance, transgenic
198 potato lines were analysed by Southern blotting and representative lines with single, double and multiple insertions were chosen from each genetic background for subsequent experiments (Fig. 1). Lines D56 and D57 of cv. Désirée had single insertions of 16D10i-2, D12 had a double insertion, and D42 showed multiple copies.
Similarly, cv. Russet Burbank lines D5 and D25 had single, D20 had double and D16 had multiple insertions of the RNAi transgene. PA99N82-4 line D53 carried a single copy of 16D10i-2, D17 and D55 had two insertions, and D2 had multiple insertions.
Northern blots indicated that a greater number of pART27(16D10i-2) transgene insertions does not necessarily lead to an increased level of 16D10i-2-specific small
RNAs (Fig. 2). Even though the expression level of 16D10i-2 small RNAs was higher in cv. Désirée lines D12 and D42 (having double and multiple insertions of the RNAi transgene, respectively) than in single insertion lines D56 and D57, such relationships were not found among some of the other lines. For example, cv. Russet Burbank line
D5 only had a single insertion of pART27(16D10i-2), but its 16D10i-2 small RNA level was considerably higher than that of other lines of the same genetic background with single, double and multiple insertions. Similar results were obtained in PA99N82-4 (Fig.
2).
At 35 DAI with M. chitwoodi isolate WAMC1, the average number of egg masses per plant in cv. Désirée lines carrying 16D10i-2 was reduced significantly by 48-59% (P
< 0.05) compared to the empty vector control (Fig. 3). Similarly, the number of egg masses in lines of cv. Russet Burbank with 16D10i-2 were lowered significantly by 37-
58% (P < 0.05) relative to empty vector controls of the same genetic background. In general, there was neither a statistical difference among the 16D10i-2-carrying lines of
199 either background, nor between the two cultivars for egg mass production. The only exception was cv. Russet Burbank line D20, which supported the smallest number of
M. chitwoodi egg masses overall, thereby leading to a statistically significant difference relative to cv. Désirée D57 (P < 0.05), the line with the greatest number of egg masses.
Advanced breeding line PA99N82-4 and its transgenic 16D10i-2 derivatives did not support M. chitwoodi WAMC1 infection, and no more than 2 egg masses plant−1 were found in this background. At 55 DAI, the average number of eggs per plant was reduced significantly by 45-60% (P < 0.05) in 16D10i-2 lines of cv. Désirée compared to the empty vector control. In cv. Russet Burbank, a significant reduction of 44-57% (P <
0.05) was observed relative to the empty vector control line. Whereas the overall level of eggs per plant was similar within and between lines of cv. Désirée and cv. Russet
Burbank, some lines were statistically different from others. For example, cv. Désirée
D56 was significantly different from D57, the lines with the lowest and highest number of eggs, respectively. In addition, cv. Russet Burbank D5, which was the most resistant line in that background, showed a statistically significant difference compared to D20 and D25. Comparisons between both cultivars indicated significant differences for the least and most resistant lines, e.g., cv. Désirée D56 vs cv. Russet Burbank D20 (all P <
0.05). No eggs were found in control and transgenic lines of PA99N82-4. Comparable results were obtained when M. chitwoodi WAMC1 infection was expressed as average number of egg masses or number of eggs per g root fresh weight, with statistically significant reductions of 29-48% and 44-55% compared to empty vector controls in cv.
Désirée, and 47-68% and 47-65% in cv. Russet Burbank, respectively (P < 0.05). Taken together, these experiments show that the 16D10i-2 RNAi transgene conferred a similar
200 level of resistance against M. chitwoodi WAMC1 in both cv. Désirée and cv. Russet
Burbank. Furthermore, 16D10i-2 did not interfere with the strong resistance against
M. chitwoodi isolate WAMC1 that is mediated by the natural resistance gene RMc1(blb) and introgressed into line PA99N82-4.
RNAI TRANSGENE 16D10I-2 INCREASES RESISTANCE AGAINST M. CHITWOODI PATHOTYPE
ROZA
To analyse whether 16D10i-2 is able to provide an increased level of resistance against M. chitwoodi pathotype Roza, which is able to overcome the RMc1(blb) gene,
PA99N82-4 and its transgenic RNAi derivatives were challenged with Roza following the same procedure. At 35 DAI the average number of egg masses per plant was reduced significantly by 32-40% (P < 0.05) compared with the empty vector control (Fig.
4). One 16D10i-2 line, D2, showed a significantly lower number of egg masses relative to the wild type, but not relative to the empty vector control at P < 0.05. Importantly, when expressed as egg masses per g root fresh weight, all PA99N82-4 16D10i-2 lines, including D2, were statistically different from both the empty vector and wild type controls, and showed a reduction of 40-47% to either baseline. At 55 DAI the average number of eggs per plant was reduced significantly by 21-29% compared to the empty vector control, and resulted in a significant reduction of 23-44% when analysed as eggs per g root fresh weight (Fig. 4). These results demonstrate that 16D10i-2 significantly reduces the formation of egg masses and the development of eggs of M. chitwoodi pathotype Roza.
201
RNAI EFFECT OF 16D10I-2 IS TRANSMITTED TO M. CHITWOODI OFFSPRING AND REDUCES ITS
PATHOGENICITY
In order to examine whether 16D10i-2-mediated resistance has an effect on the pathogenicity of M. chitwoodi offspring, nematode eggs harvested from cv. Désirée carrying either the empty vector pART27 (designated as eE29) or the RNAi construct pART27(16D10i-2) (designated as eD56) were used to inoculate empty vector (pE29) or
RNAi (pD56) plants, resulting in four possible plant-nematode combinations (see
Materials and methods). Empty vector plants inoculated with M. chitwoodi eggs harvested from cv. Désirée carrying the empty vector pART27 served as baseline for all infection assay comparisons. Importantly, at 35 DAI the average number of egg masses per plant was reduced significantly by 49% (P < 0.05) in empty vector plants inoculated with M. chitwoodi eggs harvested from cv. Désirée carrying 16D10i-2 (Fig. 5). Similarly, egg masses per plant were lowered significantly by on average 38% and 54% when potato plants expressing 16D10i-2 were inoculated with nematode eggs from empty vector and 16D10i-2 plants, respectively (P < 0.05). At 55 DAI the number of eggs per plant was reduced significantly by 50% in empty vector plants inoculated with eggs from a 16D10i-2 line. Similarly, the number of eggs per plant was significantly lower (P <
0.05) in cv. Désirée carrying 16D10i-2 inoculated with M. chitwoodi eggs from empty vector (−47%) or RNAi lines (−65%). Comparable results were obtained when the infection data were analysed as egg masses or eggs per g root fresh weight. Egg masses per g root were reduced significantly by 62, 44 and 65% for plant-nematode combinations pE29-eD56, pD56-eE29 and pD56-eD56, respectively (P < 0.05). Using g root fresh weight metrics for the same host-inoculum combinations, the number of eggs
202 was significantly lowered by 43, 30 and 56%, respectively (Fig. 5). This finding indicates that the RNAi effect of 16D10i-2 is transmitted to M. chitwoodi offspring, and significantly reduces its ability to complete its lifecycle.
To complement the infection data, the relative transcript level of the M. chitwoodi
Mc16D10L effector gene, which is targeted by the 16D10i-2 RNAi construct, was analysed by qRT-PCR and Northern blots (Fig. 6). When 16D10i-2 plants were inoculated with nematode eggs from wild type plants, the transcript level of Mc16D10L in M. chitwoodi J2 offspring as detected by qRT-PCR was reduced significantly by 38%
(P < 0.05) relative to J2 from empty vector line pE29 using the same inoculum. Using the same baseline (Mc16D10L expression level in J2 from pE29 plants inoculated with eggs from wild type plants), the relative transcript level of Mc16D10L in J2 from pE29 plants and eE29 inoculum did not differ. Importantly, the transcript level of Mc16D10L was reduced significantly by on average 58, 46 and 62% (P < 0.05) in J2 from plant- nematode combinations pE29-eD56, pD56-eE29 and pD56-eD56, respectively. This means that even in J2 from empty vector plants (pE29), which do not produce 16D10i-2 small RNAs, the transcript level of the Mc16D10L target gene was reduced by almost two-thirds if the egg inoculum was harvested from 16D10i-2 plants (eD56). Northern blots confirmed the qRT-PCR results (Fig. 6).
5. 5. DISCUSSION
Root-knot nematodes are a major problem for sustainable potato production in the Pacific Northwest of the USA. This study demonstrates that plant-mediated RNAi targeting of the putative effector gene Mc16D10L increases resistance against
203
M. chitwoodi in stable transgenic lines of potato. RNAi is emerging as a promising molecular control strategy against plant-parasitic nematodes but little is known about how to optimise the resulting resistance effect. For example, only a very limited number of studies have investigated whether changing the concentrations of small RNAs affects gene silencing in the nematode. In a previous in vitro experiment J2 of G. pallida were soaked in a ten-fold serial dilution of dsRNAs targeting the neuropeptide-coding gene flp-12 (Kimber et al., 2007). It was found that J2 migration, which served as a proxy for
RNAi efficacy, diminished strongly at a dsRNA concentration of 0.1 mg ml−1. A significant effect could still be detected at 0.1 µg ml−1, but a dsRNA dilution of ≤ 10 ng ml−1 did not impair the migratory abilities of G. pallida J2 significantly (Kimber et al.,
2007). However, siRNAs targeting the same gene could be diluted as low as 10 ng ml−1 without affecting potency to reduce J2 migration (Dalzell et al., 2010). By contrast, in vitro soaking experiments aimed at down-regulating effector gene 4E02 in Heterodera glycines J2 demonstrated that low dsRNA concentrations of 2.5 mg ml−1 were more effective at reducing the transcript level of the target gene than a higher rate of 5.0 mg ml−1 (Sukno et al., 2007).
There are only a few reports regarding stable transgenic plants transformed with an RNAi transgene to increase nematode resistance, and concentration-dependent resistance effects were not examined in these systems (for recent reviews see Li et al.,
2011; Lilley et al., 2012; Elling & Jones, 2014). In this study the relationships between
RNAi transgene copy numbers, small RNA concentrations and M. chitwoodi resistance were investigated in stable transgenic potato lines. Double or multiple insertions of
16D10i-2 do not necessarily lead to a higher level of 16D10i-2-specific small RNAs than
204 single insertions. Furthermore, no significant difference in the level of resistance measured as the amount of egg masses or eggs was detected between plants that had single, double or multiple insertions of 16D10i-2. For example, cv. Désirée lines D12 and D42, with double and multiple insertions of 16D10i-2, respectively, did not differ in their small RNA level. It is conceivable that the signal in the Northern blot was saturated, and disguised a possible small RNA concentration difference between D12 and D42. However, comparisons of other lines indicate that such was not necessarily the case. Russet Burbank lines D5 and D25 both had single transgene insertions, but showed a marked difference in their 16D10i-2-specific small RNA level. Factors other than the number of RNAi transgene insertions may have had an impact on the amount of small RNAs produced. One of the main aspects to be considered in this regard are position effects, which are a result of the random integration of transgenes into a genome (Barrell et al., 2013). Position effects frequently are reported in potato and can override the regulatory control of the promoter associated with the transgene, such that the transcript level, as well as the temporal and spatial expression patterns of the transgene are altered (Meiyalaghan et al., 2006; Barrell & Conner, 2009; Jacobs et al.,
2009; Barrell et al., 2013). It has been shown that in A. thaliana methylation of the 35S promoter can cause epigenetic silencing of transgenic RNAi constructs, leading to strongly varying levels of small RNAs between lines (Kyndt et al., 2013). Thus, it is possible that in some cases one or more 16D10i-2 RNAi transgene copies were partially or fully deactivated in similar ways, thereby resulting in double or multiple insertion lines that produce small RNAs at a level equivalent to what would be expected in single insertion lines.
205
Previous studies aimed at creating stable transgenic RNAi plants to down- regulate nematode genes made use of a single genetic background of the recipient plant species (Huang et al., 2006b; Fairbairn et al., 2007; Sindhu et al., 2009; Xue et al.,
2013; Dinh et al., 2014). Solanum sect. Petota, which consists of wild and domesticated potatoes of tuber and non-tuber-bearing species, shows an exceptionally high level of genetic diversity (Hawkes, 1990; Spooner & Hijmans, 2001; Spooner, 2009; Gavrilenko et al., 2013). In this study an attempt was made to capture some of this diversity and examine whether it has an effect on in planta RNAi-mediated resistance against
M. chitwoodi by introducing the 16D10i-2 transgene into cvs Désirée and Russet
Burbank, two different cultivars of domesticated potato (S. tuberosum ssp. tuberosum) and PA99N82-4, an advanced breeding line into which traits from the wild species
S. bulbocastanum were introgressed. Regardless of the genetic background used,
16D10i-2-mediated resistance against M. chitwoodi reached comparable levels in all lines. This result suggests that there are no genotype-specific factors that would limit the use of RNAi in a broad range of germplasm, and supports previous studies, which have demonstrated that RNAi-induced silencing of the potato vacuolar invertase and petunia flavonoid biosynthesis genes is cultivar-independent (Tsuda et al., 2004; Wu et al., 2011).
In addition to genetic diversity of host germplasm used for resistance breeding, the variability of the pathogen needs to be considered. For M. chitwoodi, a system has been developed that distinguishes two races with one pathotype each, giving four different pathogenicity types that can be differentiated based on host assays with indicator plants (Brown et al., 2009; Humphreys-Pereira & Elling, 2013, 2014).
206
M. chitwoodi pathotype Roza is able to overcome the resistance gene RMc1(blb) that has been introgressed into breeding line PA99N82-4. When 16D10i-2 was introduced into
PA99N82-4 and the resulting transgenic lines inoculated with M. chitwoodi isolate Roza, the number of egg masses and eggs per g root were reduced by about 40%. This level of resistance is slightly less than what was found for M. chitwoodi isolate WAMC1 (race
1) in 16D10i-2 lines of cv. Désirée and cv. Russet Burbank. M. chitwoodi Roza is substantially more virulent than any other known isolate, with reproductive factors that can be ten times higher in Roza compared to WAMC1 (Brown et al., 2009). In this study, Roza produced five to seven times more egg masses and eggs on wild type control plants than WAMC1, even with the same amount of inoculum.
RNAi effects are known to be inherited in C. elegans, and maintenance of the
RNAi phenotype for over 80 generations has been reported, but this phenomenon seems to be restricted to germline genes (Grishok et al., 2000; Vastenhouw et al.,
2006). In contrast, RNAi effects in Meloidogyne can be inherited even when autosomal genes that are not part of the germline are targeted. Gleason et al. (2008) provided evidence that soaking of M. javanica J2 in dsRNAs complementary to the putative Cg-1 avirulence gene results in a heritable RNAi effect that is maintained for at least five generations under selection pressure. Here it is shown that M. chitwoodi that develop on 16D10i-2 potatoes transmit the plant-mediated RNAi phenotype to offspring, and that the RNAi effect is maintained for three generations or more even in the absence of selection pressure, and reduces the pathogenicity of the nematode offspring on empty vector control plants. Subsequent inoculation of 16D10i-2 plants with M. chitwoodi in which RNAi-mediated silencing of Mc16D10L already had been initiated, did not lead to
207 a significantly increased level of resistance. This is an important finding, because it suggests that nematode infection decreases regardless of the genotype of the host once the RNAi process has been initiated through a transgenic plant. It would be interesting to investigate whether this observation translates into practical applications under field conditions. For example, one could speculate that nematode resistance can be achieved in a field with mixed genotypes, as long as at least one genotype is transgenic and induces an RNAi phenotype in the nematode. In summary, the experiments described here show that plant-mediated RNAi silencing of the putative effector gene Mc16D10L results in resistance against M. chitwoodi in diverse potato germplasm and that the RNAi effect is maintained over several generations, thereby providing resistance breeding programs with an effective new tool against this important pathogen.
5. 6. ACKNOWLEDGEMENTS
The authors thank Drs Richard Hussey and Eric Davis for the kind gift of pART27(16D10i-2) and CSIRO, Australia for providing vectors pHANNIBAL and pART27. We are grateful to Joseph Cardenas for technical assistance. This research was funded by USDA, Washington State Department of Agriculture, Washington State
Potato Commission and Idaho Potato Commission. PPNS No. 0643, Department of
Plant Pathology, College of Agricultural, Human, and Natural Resource Sciences,
Agricultural Research Center, Hatch Project No. WNP00744, Washington State
University, Pullman, WA, USA.
208
5. 7. LITERATURE CITED
Anon. (2009). Meloidogyne chitwoodi and Meloidogyne fallax. EPPO Bulletin 39, 5-17.
Anon. (2011). Potatoes 2010 summary. United States Department of Agriculture/
National Agricultural Statistics Service.
Anon. (2012). FAOSTAT. Food and Agriculture Organization of the United Nations.
Anon. (2013). Crop production. United States Department of Agriculture/National
Agricultural Statistics Service.
Barrell, P.J. & Conner, A.J. (2009). Expression of a chimeric magainin gene in potato
confers improved resistance to the phytopathogen Erwinia carotovora. Open Plant
Science Journal 3, 14-21.
Barrell, P.J., Meiyalaghan, S., Jacobs, J.M.E. & Conner, A.J. (2013). Applications of
biotechnology and genomics in potato improvement. Plant Biotechnology Journal 11,
907-920.
Brodie, B.B., Evans, K. & Franco, J. (1993). Nematode parasites of potatoes. In: Evans,
K., Trudgill, D.L. & Webster, J.M. (Eds). Plant parasitic nematodes in temperate
agriculture. Wallingford, UK, CAB International, pp. 87-132.
Brown, C.R., Mojtahedi, H. & Santo, G.S. (1989). Comparison of reproductive efficiency
of Meloidogyne chitwoodi on Solanum bulbocastanum in soil and in vitro tests. Plant
Disease 73, 957-959.
Brown, C.R., Mojtahedi, H. & Santo, G.S. (1991a). Resistance to Columbia root-knot
nematode in Solanum spp. and in hybrids of S. hougasii with tetraploid cultivated
potato. American Potato Journal 68, 445-452.
209
Brown, C.R., Yang, C.P., Kwiatkowski, S. & Adiwilaga, K.D. (1991b). Insert copy
number, chromosome-number, pollen stability, and crossability of Agrobacterium-
transformed diploid potato. American Potato Journal 68, 317-330.
Brown, C.R., Mojtahedi, H. & Santo, G.S. (1995). Introgression of resistance to
Columbia and northern root-knot nematodes from Solanum bulbocastanum into
cultivated potato. Euphytica 83, 71-78.
Brown, C.R., Mojtahedi, H. & Bamberg, J. (2004). Evaluation of Solanum fendleri as a
source of resistance to Meloidogyne chitwoodi. American Journal of Potato
Research 81, 415-419.
Brown, C.R., Mojtahedi, H., James, S., Novy, R.G. & Love, S. (2006). Development and
evaluation of potato breeding lines with introgressed resistance to Columbia root-
knot nematode (Meloidogyne chitwoodi). American Journal of Potato Research 83,
1-8.
Brown, C.R., Mojtahedi, H., Zhang, L.H. & Riga, E. (2009). Independent resistant
reactions expressed in root and tuber of potato breeding lines with introgressed
resistance to Meloidogyne chitwoodi. Phytopathology 99, 1085-1089.
Brown, C.R., Zhang, L. & Mojtahedi, H. (2013). Tracking the RMc1 gene for resistance
to race 1 of Columbia root-knot nematode (Meloidogyne chitwoodi) in three Mexican
wild potato species with different ploidies. American Journal of Potato Research 91,
180-185.
Dalzell, J.J., McMaster, S., Fleming, C.C. & Maule, A.G. (2010). Short interfering RNA-
mediated gene silencing in Globodera pallida and Meloidogyne incognita infective
stage juveniles. International Journal for Parasitology 40, 91-100.
210
Dinh, P.T.Y., Brown, C.R. & Elling, A.A. (2014). RNA interference of effector gene
Mc16D10L confers resistance against Meloidogyne chitwoodi in Arabidopsis and
potato. Phytopathology, in press.
Dropkin, V.H. (1988). The concept of race in phytonematology. Annual Review of
Phytopathology 26, 145-161.
Elling, A.A. (2013). Major emerging problems with minor Meloidogyne species.
Phytopathology 103, 1092-1102.
Elling, A.A. & Jones, J.T. (2014). Functional characterization of nematode effectors in
plants. In: Birch, P., Jones, J. & Bos, J.I.B. (Eds). Plant-pathogen interactions:
methods and protocols. Methods in Molecular Biology 1127, pp. 113-124.
Fairbairn, D.J., Cavallaro, A.S., Bernard, M., Mahalinga-Iyer, J., Graham, M.W. &
Botella, J.R. (2007). Host-derived RNAi: an effective strategy to silence genes in
plant parasitic nematodes. Planta 226, 1525-1533.
Finley, A.M. (1981). Histopathology of Meloidogyne chitwoodi (Golden et al.) on Russet
Burbank potato. Journal of Nematology 13, 486-491.
Fire, A., Xu, S., Montgomery, M.K., Kostas, S.A., Driver, S.E. & Mello, C.C. (1998).
Potent and specific genetic interference by double-stranded RNA in Caenorhabditis
elegans. Nature 391, 806-811.
Gavrilenko, T., Antonova, O., Shuvalova, A., Krylova, E., Alpatyeva, N., Spooner, D.M.
& Novikova, L. (2013). Genetic diversity and origin of cultivated potatoes based on
plastid microsatellite polymorphism. Genetic Resources and Crop Evolution 60,
1997-2015.
211
Gleason, C.A., Liu, Q.L. & Williamson, V.M. (2008). Silencing a candidate nematode
effector gene corresponding to the tomato resistance gene Mi-1 leads to acquisition
of virulence. Molecular Plant-Microbe Interactions 21, 576-585.
Grishok, A., Tabara, H. & Mello, C.C. (2000). Genetic requirements for inheritance of
RNAi in C. elegans. Science 287, 2494-2497.
Hammond, S.M., Caudy, A.A. & Hannon, G.J. (2001). Post-transcriptional gene
silencing by double-stranded RNA. Nature Reviews Genetics 2, 110-119.
Hawkes, J.G. (1990). The potato: evolution, biodiversity and genetic resource. London,
UK, Belhaven Press.
Hawkes, J.G. (1994). Origins of cultivated potatoes and species relationships. In:
Bradshaw, J.E. & Mackay, G.R. (Eds). Potato genetics. Wallingford, UK, CAB
International, pp. 3-42.
Huang, G., Dong, R., Allen, R., Davis, E.L., Baum, T.J. & Hussey, R.S. (2006a). A root-
knot nematode secretory peptide functions as a ligand for a plant transcription factor.
Molecular Plant-Microbe Interactions 19, 463-470.
Huang, G., Allen, R., Davis, E.L., Baum, T.J. & Hussey, R.S. (2006b). Engineering
broad root-knot resistance in transgenic plants by RNAi silencing of a conserved and
essential root-knot nematode parasitism gene. Proceedings of the National
Academy of Sciences of the United States of America 103, 14302-14306.
Humphreys-Pereira, D.A. & Elling, A.A. (2013). Intraspecific variability and genetic
structure in Meloidogyne chitwoodi from the USA. Nematology 15, 315-327.
Humphreys-Pereira, D.A. & Elling, A.A. (2014).Morphological variability in second-stage
juveniles and males of Meloidogyne chitwoodi. Nematology 16, 149-162.
212
Hussey, R.S. & Barker, K.R. (1973). A comparison of methods of collecting inocula of
Meloidogyne spp., including a new technique. Plant Disease Reporter 57, 1025-
1028.
Ingham, R.E., Hamm, P.B., Williams, R.E. & Swanson, W.H. (2000). Control of
Meloidogyne chitwoodi in potato with fumigant and non-fumigant nematicides.
Journal of Nematology (Suppl.) 32, 556-565.
Jacobs, J.M.E., Takla, M.F.G., Docherty, L.C., Frater, C.M., Markwick, N.P.,
Meiyalaghan, S. & Conner, A.J. (2009). Potato transformation with modified
nucleotide sequences of the cry9Aa2 gene improves resistance to potato tuber
moth. Potato Research 52, 367-378.
Janssen, G.J.W., von Norel, A., Verkerk-Bakker, B. & Janssen, R. (1995). Detecting
resistance to the root-knot nematodes Meloidogyne hapla and M. chitwoodi in potato
and wild Solanum spp. Potato Research 38, 353-362.
Janssen, G.J.W., von Norel, A., Verkerk-Bakker, B. & Janssen, R. (1996). Resistance to
Meloidogyne chitwoodi, M. fallax, and M. hapla in wild tuber-bearing Solanum spp.
Euphytica 92, 287-294.
Janssen, G.J.W., von Norel, A., Janssen, R. & Hoogendorn, J. (1997). Dominant and
additive resistance to the root-knot nematodes Meloidogyne chitwoodi and M. fallax
in Central American Solanum species. Theoretical and Applied Genetics 94, 692-
700.
Kimber, M.J., McKinney, S., McMaster, S., Day, T.A., Fleming, C.C. & Maule, A.G.
(2007). flp gene disruption in a parasitic nematode reveals motor dysfunction and
unusual neuronal sensitivity to RNA interference. FASEB Journal 21, 1233-1243.
213
Kyndt, T., Ji, H., Vanholme, B.&Gheysen, G. (2013). Transcriptional silencing of RNAi
constructs against nematode genes in Arabidopsis. Nematology 15, 519-528.
Li, J., Todd, T.C., Lee, J. & Trick, H.N. (2011). Biotechnological application of functional
genomics towards plant-parasitic nematode control. Plant Biotechnology Journal 9,
936-944.
Li, X.Q.,Wei, J.Z., Tan, A. & Aroian, R.V. (2007). Resistance to root-knot nematode in
tomato roots expressing a nematicidal Bacillus thuringiensis crystal protein. Plant
Biotechnology Journal 5, 455-464.
Lilley, C.J., Davies, L.J. & Urwin, P.E. (2012). RNA interference in plant parasitic
nematodes: a summary of the current status. Parasitology 139, 630-640.
Livak, K.J. & Schmittgen, T.D. (2001). Analysis of relative gene expression data using
real-time quantitative PCR and the 2–ΔΔCt method. Methods 25, 402-408.
Meiyalaghan, S., Jacobs, J.M.E., Butler, R.C., Wratten, S.D. & Conner, A.J. (2006).
Expression of cry1Ac9 and cry9Aa2 genes under a potato light-inducible Lhca3
promoter in transgenic potatoes for tuber moth resistance. Euphytica 147, 297-309.
Mitchum, M.G., Wang, X. & Davis, E.L. (2008). Diverse and conserved roles of CLE
peptides. Current Opinion in Plant Biology 11, 75-81.
Mitchum, M.G., Hussey, R.S., Baum, T.J., Wang, X., Elling, A.A., Wubben, M. & Davis,
E.L. (2013). Nematode effector proteins: an emerging paradigm of parasitism. New
Phytologist 199, 879-894.
Perry, R.N., Moens, M. & Starr, J.L. (Eds) (2009). Root-knot nematodes. Wallingford,
UK, CAB International.
214
Porter, I., Banks, J., Mattner, S. & Fraser, P. (2009). Global phaseout of methyl bromide
under the Montreal Protocol: implications for bioprotection, biosecurity and the
ozone layer. In: Gisi, U., Chet, I. & Gullino, M.L. (Eds). Recent developments in
management of plant diseases. Plant pathology in the 21st century. Berlin,
Germany, Springer-Verlag, pp. 293-309.
Sheerman, S. & Bevan, M.W. (1988). A rapid transformation method for Solanum
tuberosum using binary Agrobacterium tumefaciens vectors. Plant Cell Reports 7,
13-16.
Sindhu, A.S., Maier, T.R., Mitchum, M.G., Hussey, R.S., Davis, E.L. & Baum, T.J.
(2009). Effective and specific in planta RNAi in cyst nematodes: expression
interference of four parasitism genes reduces parasitic success. Journal of
Experimental Botany 60, 315-324.
Spooner, D.M. (2009). DNA barcoding will frequently fail in complicated groups: an
example in wild potatoes. American Journal of Botany 96, 1177-1189.
Spooner, D.M. & Hijmans, R.J. (2001). Potato systematics and germplasm collecting,
1989-2000. American Journal of Potato Research 78, 237-268.
Sukno, S.A.,McCuiston, J.,Wong, M.Y.,Wang, X., Thon, M.R., Hussey, R.S., Baum, T.J.
& Davis, E.L. (2007). Quantitative detection of double-stranded RNA-mediated gene
silencing of parasitism genes in Heterodera glycines. Journal of Nematology 39,
145-152.
Tsuda, S., Fukui, Y., Nakamura, N., Katsumoto, Y., Yonekura-Sakakibara, K., Fukuchi-
Mizutani, M., Ohira, K., Ueyama, Y., Ohkawa, H., Holton, T.A. et al. (2004). Flower
215
color modification of Petunia hybrida commercial varieties by metabolic engineering.
Plant Biotechnology 21, 377-386.
Urwin, P.E., Møller, S.G., Lilley, C.J., McPherson, M.J. & Atkinson, H. (1997). Continual
green-fluorescent protein monitoring of cauliflower mosaic virus 35S promoter
activity in nematode-induced feeding cells in Arabidopsis thaliana. Molecular Plant-
Microbe Interactions 10, 394-400.
Vastenhouw, N.L., Brunschwig, K., Okihara, K.L., Müller, F., Tijsterman, M. & Plasterk,
R.H.A. (2006). Long-term gene silencing by RNAi. Nature 442, 882.
Wu, L., Bhaskar, P.B., Busse, J.S., Zhang, R., Bethke, P.C. & Jiang, J. (2011).
Developing cold-chipping potato varieties by silencing the vacuolar invertase gene.
Crop Science 51, 981-990.
Xue, B., Hamamouch, N., Li, C., Huang, G., Hussey, R.S., Baum, T.J. & Davis, E.L.
(2013). The 8D05 parasitism gene of Meloidogyne incognita is required for
successful infection of host roots. Phytopathology 103, 175-181.
Yang, Y., Jittayasothorn, Y., Chronis, D., Wang, X., Cousins, P. & Zhong, G.Y. (2013).
Molecular characteristics and efficacy of 16D10 siRNAs in inhibiting root-knot
nematode infection in transgenic grape hairy roots. PLoS ONE 8(7), e69463.
Zhang, F., Peng, D., Ye, X., Yu, Z., Hu, Z., Ruan, L. & Sun, M. (2012). In vitro uptake of
140 kDa Bacillus thuringiensis nematicidal crystal proteins by the second stage
juvenile of Meloidogyne hapla. PLoS ONE 7(6), e38534.
216
5. 8. TABLE
Table 1. Primers and probes used for PCR, Southern and northern blots.
Probe/gene Primer name Primer sequences (5’3’)
16D10 16D10F5 GTTTACTAATTCAATTAAAAATTTAATT
16D10R5 CAATTATTTCCTCCAGGATTTGGCCC
U6 (X60506) U6F GCGCAAGGATGACACGCA
U6R GGCTGAGTTATTTTTTTCTG
ITS2 (JN241865) McITS-RTF GGGGTCAAACCCTTTGGCACGTCTGG
McITS-RTR GCGGGTGATCTCGACTGAGTTCAGG
Mc16D10L Mc16D10-F GATATTTAAATTAAATTATATTCTTCTAAA
(KF734590) Mc16D10-R GCTTTATTCAATTTATTTTTATTTATT
Mc16D10-RTF TTATTTTATCTGTTACTTTTGTGGATTCAGC
Mc16D10-RTR GCGACCATCATTATTATCATTTCCACC
16D10i-2 35S-F TTCGCAAGACCCTTCCTCTA
OCS1 CTTCTTCGTCTTACACATCACTTGTC
nptII-F ATCGGGAGCGGCGATACCGTA
nptII-R GACGCTATTCGGCTATGACTG
217
5. 9. FIGURES
Fig. 1. Southern blots showing copy numbers of 16D10i-2 in stable transgenic potato lines. Lines based on cv. Désirée include DES, wild type; E29, empty vector pART27;
D56, D57, D12, D42, transformed with pART27(16D10i-2). Lines based on cv. Russet
Burbank include RB, wild type; E34, empty vector pART27; D5, D25, D20, D16, transformed with pART27(16D10i-2). Lines based on advanced breeding line PA99N82-
4 include 82-4, wild type; E12, empty vector pART27; D17, D53, D55, D2, transformed with pART27(16D10i-2). Probe 16D10i-2 (Table 1) was used for the hybridisation.
218
Fig. 2. Northern blots for stable transgenic potato lines. A: 16D10i-2-specific small
RNAs hybridising with probe 16D10; B: U6 small nuclear RNA (snRNA) loading control;
C: Total RNA loading control. DES, wild type; E29, empty vector pART27; D56, D57,
D12, D42, transformed with pART27(16D10i-2), all with cv. Désirée as genetic background. RB, wild type; E34, empty vector pART27; D5, D25, D20, D16, transformed with pART27(16D10i-2), all with cv. Russet Burbank as genetic background. 82-4, wild type; E12, empty vector pART27; D17, D53, D55, D2, transformed with pART27(16D10i-2), all with advanced breeding line PA99N82-4 as genetic background.
219
Fig. 3. Reproductive success of Meloidogyne chitwoodi WAMC1 on transgenic potato lines expressing 16D10i-2. A: Number of egg masses per plant at 35 days after inoculation (DAI); B: Number of egg masses per g root fresh weight at 35 DAI; C:
Number of eggs per plant at 55 DAI; D: Number of eggs per g root fresh weight at 55
DAI. DES, wild type; E29, empty vector pART27; D56, D57, D12, D42, transformed with pART27(16D10i-2), all with cv. Désirée as genetic background. RB, wild type; E34, empty vector pART27; D5, D25, D20, D16, transformed with pART27(16D10i-2), all with cv. Russet Burbank as genetic background. 82-4, wild type; E12, empty vector pART27;
D17, D53, D55, D2, transformed with pART27(16D10i-2), all with advanced breeding line PA99N82-4 as genetic background. Each bar represents the mean of ten plants per independent line and time-point with standard errors. Letters indicate statistically significant differences using a Student’s t -test (P < 0.05).
220
Fig. 4. Reproductive success of Meloidogyne chitwoodi Roza on transgenic potato lines expressing 16D10i-2 in a PA99N82-4 genetic background. A: Number of egg masses per plant at 35 days after inoculation (DAI); B: Number of egg masses per g root fresh weight at 35 DAI; C: Number of eggs per plant at 55 DAI; D: Number of eggs per g root fresh weight at 55 DAI. 82-4, wild type; E12, empty vector pART27; D17, D53, D55, D2, transformed with pART27(16D10i-2), all with advanced breeding line PA99N82-4 as genetic background. Each bar represents the mean of ten plants per independent line and time-point with standard errors. Letters indicate statistically significant differences using a Student’s t -test (P < 0.05).
221
Fig. 5. Pathogenicity and reproductive success of M. chitwoodi offspring from potato lines with and without the 16D10i-2 RNAi transgene. A: Number of egg masses per plant at 35 days after inoculation (DAI). B: Number of egg masses per g root fresh weight at 35 DAI; C: Number of eggs per plant at 55 DAI; D: Number of eggs per g root fresh weight at 55 DAI. M. chitwoodi WAMC1 inoculum was harvested from transgenic potato cv. Désirée lines E29 (empty vector control) and D56 (carrying pART27(16D10i-
2)), resulting in four different treatments: empty vector potato line E29 inoculated with
M. chitwoodi eggs collected from line E29 (pE29-eE29); empty vector line E29 inoculated with M. chitwoodi eggs collected from 16D10i-2 line D56 (pE29-eD56),
16D10i-2 line D56 inoculated with M. chitwoodi eggs collected from line empty vector line E29 (pD56-eE29) and 16D10i-2 line D56 inoculated with M. chitwoodi eggs collected from line D56 (pD56-eD56), where ‘p’ stands for plant and ‘e’ stands for eggs.
M. chitwoodi isolate used was WAMC1. Each bar represents the mean of ten plants per
222 independent line and time-point with standard errors. Letters indicate statistically significant differences using a Student’s t -test (P < 0.05).
223
Fig. 6. Relative fold change of Mc16D10L transcript level in Meloidogyne chitwoodi second-stage juveniles (J2) offspring from different potato-nematode combinations. J2 were hatched from eggs collected from infection assays pE29, pD56, pE29-eE29, pE29- eD56, pD56-eE29 and pD56-eD56. A: qRT-PCR showing relative fold changes of
Mc16D10L transcript level in J2. Mc16D10L expression level in J2 from eggs harvested from pE29 was set as 1. Each bar represents the mean of qRT-PCR reactions run in triplicate with standard errors. Letters indicate statistically significant differences using a
Student’s t - test (P < 0.05); B: Northern blot showing Mc16D10L transcript level in J2 using probe Mc16D10L (Table 1); C: Northern blot for ITS2 expression as control; D:
Agarose gel showing total RNA loading control. Genetic background of all potato lines used was cv. Désirée and M. chitwoodi isolate was WAMC1. pE29, empty vector line
E29 inoculated with M. chitwoodi collected from wild type tomatoes cv. Rutgers; pD56,
16D10i-2 line D56 inoculated with M. chitwoodi collected from wild type tomatoes cv.
Rutgers; pE29-eE29, empty vector line E29 inoculated with M. chitwoodi eggs collected from line E29; pE29-eD56, empty vector line E29 inoculated with M. chitwoodi eggs collected from 16D10i-2 line D56; pD56-eE29, 16D10i-2 line D56 inoculated with
M. chitwoodi eggs collected from line empty vector line E29; pD56-eD56, 16D10i-2 line
D56 inoculated with M. chitwoodi eggs collected from line D56, where ‘p’ stands for plant and ‘e’ stands for eggs. Eggs harvested to hatch J2 for qRT-PCR and Northern
224 blots are from the same plants analysed for pathogenicity of M. chitwoodi offspring as shown in Figure 5.
225
CHAPTER SIX
BROAD MELOIDOGYNE RESISTANCE IN POTATO BASED ON RNA
INTERFERENCE OF EFFECTOR GENE 16D10
Phuong T.Y. Dinh1, Linhai Zhang2, Hassan Mojtahedi3, Charles R. Brown2,
and Axel A. Elling1
1Department of Plant Pathology, Washington State University, Pullman, WA 99164.
2Vegetable and Forage Crops Research Unit, United States Department of Agriculture –
Agricultural Research Service, Prosser, WA 99350.
3Washington State Department of Agriculture, Prosser, WA 99350.
Corresponding author : Axel A. Elling
E-mail: [email protected]
This paper was published in 2015 in Journal of Nematology 47(1):71-78
Running head: Broad Meloidogyne RNAi resistance in potato: Dinh et al.
Key words: Effector, host-parasitic relationship, Meloidogyne arenaria, Meloidogyne chitwoodi, Meloidogyne hapla, Meloidogyne incognita, Meloidogyne javanica, potato
(Solanum tuberosum), resistance, RNA interference.
This paper was edited by Andrea Skantar.
226
6. 1. ABSTRACT
Root-knot nematodes (Meloidogyne spp.) are a significant problem in potato
(Solanum tuberosum) production. There is no potato cultivar with Meloidogyne resistance, even though resistance genes have been identified in wild potato species and were introgressed into breeding lines. The objectives of this study were to generate stable transgenic potato lines in a cv. Russet Burbank background that carry an RNA interference (RNAi) transgene capable of silencing the 16D10 Meloidogyne effector gene, and test for resistance against some of the most important root-knot nematode species affecting potato, i.e., M. arenaria, M. chitwoodi, M. hapla, M. incognita, and
M. javanica. At 35 days after inoculation (DAI), the number of egg masses per plant was significantly reduced by 65% to 97% (P < 0.05) in the RNAi line compared to wild type and empty vector controls. The largest reduction was observed in M. hapla, whereas the smallest reduction occurred in M. javanica. Likewise, the number of eggs per plant was significantly reduced by 66% to 87% in M. arenaria and M. hapla, respectively, compared to wild type and empty vector controls (P < 0.05). Plant-mediated RNAi silencing of the 16D10 effector gene resulted in significant resistance against all of the root-knot nematode species tested, whereas RMc1(blb), the only known Meloidogyne resistance gene in potato, did not have a broad resistance effect. Silencing of 16D10 did not interfere with the attraction of M. incognita second-stage juveniles to roots, nor did it reduce root invasion.
227
6. 2. INTRODUCTION
Root-knot nematodes (Meloidogyne spp.) pose serious problems to potato
(Solanum tuberosum) production in both temperate and tropical climates (Brodie et al.,
1993; Sikora and Fernández, 2005). In addition to direct losses resulting from reduced tuber yields, Meloidogyne spp. can render potatoes unmarketable because of quality defects caused by gall formation on the tuber surface (Finley, 1981). Some species are quarantine pests and interfere with international trade. Even though M. chitwoodi is present in most major potato-producing areas, its detection in tubers can lead to the rejection of entire shipments (Ingham et al., 2000; Elling, 2013). To date, no commercially available potato cultivars with resistance to root-knot nematodes exist.
Current nematode control strategies in potato heavily rely on synthetic nematicides.
This practice is not only costly but also potentially harmful to the environment and faces increasing restrictions from regulatory agencies in many countries.
Knowledge about root-knot nematode resistance genes in potato is sparse
(Gebhardt and Valkonen, 2001; Sanchez-Puerta and Masuelli, 2011). There is no known root-knot nematode resistance gene in the cultivated potato S. tuberosum, but several genes have been found in wild potato species (Brown et al., 1991a, 1995;
Janssen et al., 1996; Brown et al., 2004; Williamson and Kumar, 2006). The best characterized root-knot nematode resistance gene in Solanum sect. Petota is RMc1(blb) from S. bulbocastanum, a gene that is effective against some races of M. chitwoodi
(Brown et al., 2009). The resistance mechanism of RMc1(blb) is based on a hypersensitive response and involves calcium signaling (Davies et al., 2015). Recent studies suggest that M. chitwoodi resistance in different species of Solanum is based on the same gene,
228 thereby limiting the diversity of available resistance (Brown et al., 2014).
Resistance based on plant-mediated RNA interference (RNAi) is emerging as a promising new disease control tactic. RNAi was first discovered in Caenorhabditis elegans but is now regarded as a widespread phenomenon in virtually all eukaryotes
(Fire et al., 1998). RNAi is based on the ability of a cell to detect and degrade double- stranded RNA (dsRNA). Dicer, a ribosome III-like enzyme, catalyzes the cleavage of long dsRNA into small interfering RNA (siRNA) segments of about 21 nt in length.
Double-stranded siRNA molecules are unwound and separated into single strands.
While the passenger strand is degraded, the guide strand is loaded into a large complex, RISC (RNA-induced silencing complex). If the guide strand binds to a complementary region in the mRNA of a target gene, the respective gene is silenced
(Hammond et al., 2001).
Endogenous nematode genes can be silenced by expressing dsRNA that is complementary to nematode genes in planta. Since root-knot nematodes are able to ingest molecules of up to 140 kDa (Zhang et al., 2012), there is an opportunity for oral delivery of dsRNA or siRNA into the nematode. During feeding, the nematodes take up plant cytoplasm containing dsRNA or siRNA that may then silence the respective target gene(s) in the parasite (Lilley et al., 2012; Elling and Jones, 2014). For RNAi applications in an agricultural setting, it is essential to avoid off-target effects that could silence genes in the host plant or nontarget animals or humans. Nematode effector genes represent a very specific and therefore attractive target. Effector genes are expressed in the esophageal gland cells of plant-parasitic nematodes and form the basis for the molecular interactions between the parasite and its host (Mitchum et al.,
229
2013). The 16D10 effector gene was initially cloned in M. incognita, but orthologs have since been identified in M. arenaria, M. chitwoodi, M. hapla, and M. javanica (Huang et al., 2006a, 2006b; Dinh et al., 2014a). The nucleotide sequence of 16D10 is 95 to 98% identical between M. arenaria, M. hapla, M. incognita, and M. javanica (Huang et al.,
2006a). In contrast, the M. chitwoodi ortholog is only 70% identical to M. incognita (Dinh et al., 2014a). Previous studies have shown that plant-mediated silencing of the
Meloidogyne effector gene 16D10 can lead to a dramatic increase in resistance (Huang et al., 2006a; Dinh et al., 2014a, 2014b). However, these reports were based either on
Arabidopsis or made use of only a single species of Meloidogyne. The objective of this study was to select a high-performing stable transgenic 16D10 RNAi line with the genetic background of a commercial potato cultivar and then evaluate the resistance of this line against a broad range of Meloidogyne spp. found in temperate and tropical potato-producing regions.
6. 3. MATERIALS AND METHODS
Plant transformation: Stable transgenic lines of potato ‘Russet Burbank’ were generated as described by Dinh et al. (2014a). Briefly, potato stem segments (about 1 cm in length) obtained from plants growing under axenic conditions were coincubated with Agrobacterium tumefaciens strain GV3101 carrying silencing vector pART27(16D10i-2) (Huang et al., 2006a) or empty vector pART27 as control. Incubation was done for 3 d in the dark at 19°C on CIM media (MS basal salts, 0.25 ppm folic acid,
0.05 ppm D-biotin, 2 ppm glycine, 0.5 ppm nicotinic acid, 0.5 ppm pyridoxine HCl, 0.4 ppm thiamine HCl, 0.01% myo-inositol, 3% D-sucrose, 1 ppm 6-benzylaminopurine, 2
230 ppm 1-naphthaleneacetic acid, 0.6% Daishin agar, pH 5.6). The stem segments were transferred every 2 wk for up to 3 mon to fresh 3C5ZR media (MS basal salts, 0.5 ppm nicotinic acid, 0.5 ppm pyridoxine HCl, 1 ppm thiamine HCl, 0.01% myo-inositol, 3% D- sucrose, 0.5 ppm indole-3-acetic acid, 3 ppm zeatin ribose, 0.5 g/liter timentin, 70 µg/ml kanamycin sulfate, 0.6% Daishin agar, pH 5.9), supplemented with 50 µg/ml kanamycin sulfate and 50 µg/ml timentin (Sheerman and Bevan, 1988; Brown et al., 1991b; Dinh et al., 2014a). Potato plantlets that regenerated on 3C5ZR media were maintained and propagated on propagation media (MS basal salts, 3% D-sucrose, 50 µg/ml kanamycin sulfate, 50 µg/ml timentin, 0.6% Daishin agar, pH 5.7). Wild type ‘Russet Burbank’ and advanced breeding line PA99N82-4 were maintained on propagation media without kanamycin sulfate and timentin. PA99N82-4 carries the RMc1(blb) resistance gene from
S. bulbocastanum (Brown et al., 2009).
Southern and northern blotting: DNA was extracted from potato leaves ground in liquid nitrogen with TPS buffer (100 mM Tris-HCl pH 8, 100 mM EDTA pH 8, 1 M KCl), precipitated with isopropanol, treated with RNase, and purified with chloroform/isopropanol precipitation. The DNA pellets were resuspended in 100 µl sterile water before being used for Southern blots. For each plant line, 15 µg DNA was digested with 50 U Xba I (New England Biolabs, Ipswich, MA) for 16 hr at 37°C, then separated on a 0.8% agarose gel at 70 V for 16 hr and transferred to a GeneScreen
Plus nylon membrane (PerkinElmer, Boston, MA) in 10× saline sodium citrate (SSC) buffer (1.5 M NaCl, 0.15 M sodium citrate, pH 7). The membrane was UV cross-linked and hybridized for 16 hr at 42°C with a [α-32P] dATP labeled 16D10i-2 probe (Dinh et al., 2014b) in hybridization buffer (50% deionized formamide, 0.1 mg/ml salmon sperm
231
DNA, 1% sodium dodecyl sulfate (SDS), 1 M NaCl, 10% dextran sulfate). Following hybridization, the membrane was washed twice with 2× SSC buffer for 5 min at 42°C, three times with 2× SSC plus 1% SDS for 20 min at 65°C, and three times with 0.1×
SSC plus 1% SDS for 20 min at 42°C. After washing, the membrane was exposed to X- ray film for 2 d at –70°C.
For northern blots, total RNA enriched for small RNA was extracted from 1 g potato leaves using the mirVana miRNA isolation kit (Ambion, Austin, TX) according to the manufacturer’s instructions. Twenty micrograms denatured total RNA of each potato line was separated on a 1% agarose gel at 120 V for 1 hr, transferred to a Nytran N nylon membrane (Sigma-Aldrich, St. Louis, MO) for 16 hr, UV cross-linked, and hybridized with [α-32P] dATP labeled 16D10 and U6 probes (Dinh et al., 2014b). After 16 hr of hybridization at 25°C in hybridization buffer (50% deionized formamide, 3× SSC,
0.1 mg/ml salmon sperm DNA, 1% SDS, 0.05 M phosphate buffer, 0.2% bovine serum albumin, 0.2% polyvinylpyrrolidone, 0.2% Ficoll), the membrane was washed three times for 20 min at 46°C with 2× SSC plus 0.2% SDS, and exposed to X-ray film for 1 to
5 d at –70°C. Details about probes used for Southern and northern blots were described previously by Dinh et al. (2014a, 2014b).
Nematode culture and extraction: Meloidogyne arenaria, M. chitwoodi isolate
WAMC1, M. hapla, M. incognita isolate OP-50, and M. javanica were maintained on tomato (S. lycopersicum) ‘Rutgers’ grown in autoclaved sand under greenhouse conditions. Nematode eggs were extracted from roots 3 mon postinoculation. To release the eggs, roots were cut into 1- to 3-cm pieces and agitated in a 0.5% NaOCl solution following the method of Hussey and Barker (1973). The suspension was
232 poured over nested sieves (850, 75, 25 µm pore size from top to bottom) and eggs collected on the 25 µm pore size sieve. Eggs were purified on a 70% sucrose gradient
(Elling and Jones, 2014) before being used for infection assays. To obtain M. incognita infective second stage juveniles (J2) for attraction assays, eggs were hatched in modified Baermann pans (Dinh et al., 2014a).
Nematode infection assays: Single nodes of each potato line were grown in propagation media for 30 d before being transferred to a sterilized sand/soil mixture (3 parts sand : 1 part soil). Plants were supplied with 20-10-20 N-P-K liquid fertilizer every
2 d for the duration of the experiment. All infection assays were set up in randomized complete block designs with host genotype as the main effect.
The initial infection assay screened for highly resistant potato lines. Five plants each of 22 transgenic 16D10i-2 lines and wild type ‘Russet Burbank’ were grown in separate 6-in clay pots and inoculated with 1,200 M. chitwoodi eggs per pot after the plants were acclimated to greenhouse conditions (16-hr light, 22°C and 8-hr dark, 20°C) for 10 d. Roots were harvested at 55 days after inoculation (DAI), washed to remove soil particles, and processed for egg extraction as described above. The total number of eggs was estimated by counting three 100-µl aliquots of a 50-ml suspension per root system under a Stemi 2000C stereomicroscope (Zeiss, Jena, Germany). The reproductive efficiency R was calculated as the ratio between the initial inoculated number of eggs (Pi) and the final number of eggs at harvest (Pf).
For subsequent infection assays, the best performing transgenic line D21 was tested for resistance against M. arenaria, M. hapla, M. incognita, and M. javanica, with empty vector line E34, wild type PA99N82-4, and ‘Russet Burbank’ controls. Plantlets
233 were transferred to Ray Leach SC10U cone-tainers (Stuewe and Sons, Tangent, OR) and acclimated to a growth chamber for 10 d before being inoculated with nematode eggs. The infection assay was performed under 16-hr light, 24°C and 8-hr dark, 22°C conditions in a growth chamber. For each line, 20 plants were inoculated with 2,000 eggs per nematode species. At 35 DAI, 10 root systems were harvested to count egg masses and at 55 DAI an additional 10 root systems were harvested to count the number of eggs, respectively, for each plant line and nematode combination. At 35 DAI, roots were washed, fresh weight per root system determined, and roots were stained for
15 min with 0.15 g/liter phloxine B (Fisher Scientific, Fair Lawn, NJ) to visualize egg masses. At 55 DAI, root fresh weight was determined and eggs extracted as described above. The infection assay was conducted twice for each plant line and nematode combination.
Nematode attraction assays: The attraction of M. incognita J2 to 16D10i-2 potato line D21 and PA99N82-4, empty vector E34 and wild type ‘Russet Burbank’ controls was investigated following the methods of Wang et al. (2009). Briefly, potato plantlets grown for 15 d from single nodes in propagation media were placed on microscope glass slides. The root systems were coated with 1 ml of 23% pluronic gel PF-127
(Sigma-Aldrich) containing 300 M. incognita J2. For each potato line, 15 replicates with one plantlet each were prepared on separate glass slides and then covered with cover slips. After exposure in a moist chamber for 6 hr at 25°C, the numbers of J2 touching the terminal 10 mm of each root tip were counted. Following, samples were kept in a moist chamber for an additional 4 d before the roots were stained with acid fuchsin
(Acros Organics, Morris Plains, NJ). For staining, roots were soaked in 0.5% NaOCl for
234
3 min, washed under running tap water for 10 min, and boiled in acid fuchsin solution
(0.35% acid fuchsin, 25% acetic acid) for 5 min (Byrd et al., 1983). Stained roots were kept in distilled water at 4°C until parasitic J2 inside roots were counted. Nematode attraction and invasion were visualized with a SteREO Discovery.V8 stereomicroscope,
AxioCam ICc1 digital camera and ZEN imaging software (Zeiss) at 6 h after inoculation and 4 DAI and staining, respectively.
Data analysis: Average numbers of eggs and egg masses and SE were calculated using Microsoft Excel. A Student’s t-test (LSD) at alpha level 0.05 (SAS 9.2 software; SAS Institute, Cary, NC) was used to estimate statistical significant differences between treatments.
6. 4. RESULTS
Screening of 16D10i-2 stable transgenic potato lines for resistance against
M. chitwoodi: Silencing construct pART27(16D10i-2) was introduced into ‘Russet
Burbank’ potato to generate stable transgenic RNAi lines. Twenty-two independent lines, none of which showed any overt phenotypical changes compared to wild type plants, were chosen for further analysis. Southern blots indicated that lines D3, D5,
D11, D21, D24 and D25 carried single copies of the 16D10i-2 transgene. Lines D2, D16 and D20 had double insertions, and the remaining lines carried more than two copies of
16D10i-2 (Fig. 1). For the 22 transgenic RNAi lines and wild type control plants inoculated with M. chitwoodi eggs, the number of eggs at 55 DAI and Pf/Pi ratios revealed that lines D1, D9, D21, D25, D33, and D36 were most resistant against
M. chitwoodi. The average number of eggs per plant at 55 DAI in the resistant lines
235 ranged from 263 in D21 to 5,633 in D25, and had corresponding Pf/Pi ratios from 0.22 to 4.69. The reduction was significant (P < 0.05) compared to wild type plants, which yielded on average 98,882 eggs per plant and had a Pf/Pi ratio of 82.4 (Fig. 2, Table 1).
D21 emerged as the most highly resistant line in this screening assay, and was chosen for further analysis. Northern blots confirmed a strong expression of 16D10-specific small RNAs in line D21 (Fig. 3).
Broad Meloidogyne resistance in stable transgenic 16D10i-2 potato line D21:
When challenged with M. arenaria, M. hapla, M. incognita, or M. javanica, line D21 showed strong resistance regardless of root-knot nematode species. At 35 DAI, the number of egg masses per plant was significantly reduced (P < 0.05) in D21 compared to wild type ‘Russet Burbank’, empty vector and PA99N82-4 control plants by 65 to 97%
(Fig. 4,5). The greatest reduction in the number of egg masses was achieved against
M. hapla, whereas M. javanica showed the smallest reduction. Comparable levels of resistance were observed when the number of egg masses was analyzed on a per gram root basis (Fig. 4A,B). PA99N82-4 control plants, which carried the RMc1(blb) resistance gene, had significantly fewer egg masses for M. arenaria and M. incognita on a per plant basis, but not when the data were adjusted for root fresh weight (P < 0.05).
At 55 DAI, the number of eggs per plant remained significantly lower in line D21 for all root-knot nematodes tested (P < 0.05). Compared to wild type and empty vector controls, egg production in line D21 was reduced from 66% in M. arenaria to 87% in
M. hapla on a per plant basis (Fig. 4C). Similar levels of resistance were achieved when the number of eggs were expressed on a per gram root basis (Fig. 4D). Line PA99N82-
236
4 significantly lowered the number of eggs on a per plant and per gram root basis for
M. hapla, but not for the other root-knot nematodes tested.
Plant-mediated 16D10i-2 resistance does not affect Meloidogyne attraction to host roots: To test whether 16D10i-2 alters the nematodes’ ability to find and invade roots, an attraction assay was performed. There was no statistically significant difference between D21 and PA99N82-4, ‘Russet Burbank’ and empty vector controls when the number of M. incognita infective J2 or parasitic J2 were counted at 6 hr after inoculation or 4 DAI, respectively. No overt phenotypical differences were observed between the roots of either line (Fig. 6).
6. 5. DISCUSSION
This study demonstrates that plant-mediated RNAi resistance can protect potato against important root-knot nematode pathogens. Stable transgenic lines of potato expressing the RNAi transgene 16D10i-2 showed significant resistance against
M. arenaria, M. chitwoodi, M. hapla, M. incognita, and M. javanica. The 16D10 gene is widely conserved in Meloidogyne spp., but sequence divergence can reach about 30% between some species. Importantly, all orthologs found so far carry a highly conserved region of about 21 nt that is targeted by the 16D10i-2 silencing construct (Dinh et al.,
2014a). Thus, statistically significant increases in resistance compared to wild type and empty vector controls could be achieved for all root-knot nematode species tested here.
The number of RNAi transgene insertions into the host genome was not a reliable indicator of the resistance level. This finding confirms previous results and underscores the dominating role of position effects (Barrell et al., 2013; Dinh et al., 2014b).
237
Furthermore, one or more copies of promoters used to control exogenous silencing genes targeted at nematodes can be affected by epigenetic silencing in the plant. This phenomenon has recently been demonstrated for the Cauliflower mosaic virus 35S promoter (Kyndt et al., 2013), a ubiquitous promoter that also was used to control
16D10i-2 in this study.
Breeding for resistance against pathogens is a high priority in integrated pest management strategies. Unfortunately, the number of known root-knot nematode resistance genes in potato is extremely limited (Gebhardt and Valkonen, 2001;
Williamson and Kumar, 2006). Previous studies indicated that moving root-knot nematode resistance genes across species barriers within the Solanaceae can lead to resistance in the recipient plant (Goggin et al., 2006). However, this approach is limited by the fact that the respective resistance gene might not be effective against root-knot nematodes that are relevant in the recipient plant species (Brown et al., 1997). As an alternative, this study demonstrates that targeting effector genes that are highly conserved between a wide range of Meloidogyne spp. may be a promising control strategy and should lead to significant increases in nematode resistance.
Even though RNAi-based resistance is able to suppress the reproductive success of the nematode, this approach does not lend itself to shielding the plant from the primary infection. For a plant-mediated RNAi strategy to work, the nematode must take up plant cytoplasm containing small RNA or dsRNA complementary to an endogenous gene. Therefore, only feeding nematode life stages are vulnerable to acquiring the silencing trigger from the host. Non-feeding life stages, such as the infective J2 are free to invade host roots, however. In M. chitwoodi, the 16D10 transcript
238 is highly upregulated in infective J2, and in M. incognita antiserum confirmed the accumulation of 16D10 peptide in the subventral gland cells (Huang et al., 2006b; Dinh et al., 2014a). In this study, no difference was detected in the ability of M. incognita J2 to locate or invade roots of potato plants expressing or lacking the silencing gene
16D10i-2. This observation suggests that 16D10i-2-mediated resistance mechanisms occur after nematodes have invaded host roots and become sedentary. Importantly, the
16D10i-2 silencing effect can be transmitted epigenetically to nematode offspring and provide resistance even in non-RNAi plants (Dinh et al., 2014b).
In summary, this study demonstrates that RNAi-induced silencing of a nematode effector gene can lead to a dramatic reduction of a broad range of Meloidogyne spp. in stable transgenic lines of ‘Russet Burbank’ potato, thereby providing an attractive new tool for molecular breeding strategies against root-knot nematodes in this important crop.
6. 6. ACKNOWLEDGMENTS
The authors thank Dr. Debra Inglis for critical reading of an earlier version of the manuscript. Funding for this project was provided by U.S. Department of Agriculture,
Washington State Department of Agriculture, and Northwest Potato Research
Consortium. PPNS No. 0672, Department of Plant Pathology, College of Agricultural,
Human, and Natural Resource Sciences, Agricultural Research Center, Hatch Project
No. WNP00744, Washington State University, Pullman, WA 99164.
239
6. 7. LITERATURE CITED
Barrell, P. J., Meiyalaghan, S., Jacobs, J. M. E., and Conner, A. J. 2013.
Applications of biotechnology and genomics in potato improvement. Plant
Biotechnology Journal 11:907–920.
Brodie, B. B., Evans, K., and Franco, J. 1993. Nematode parasites of potatoes.
Pp. 87–132 in K. Evans, D. L. Trudgill, J. M. Webster, eds. Plant parasitic nematodes in temperate agriculture. Wallingford, UK: CAB International.
Brown, C. R., Mojtahedi, H., and Santo, G. S. 1991a. Resistance to Columbia root-knot nematode in Solanum spp. and in hybrids of S. hougasii with tetraploid cultivated potato. American Journal of Potato Research 68:445–452.
Brown, C. R., Mojtahedi, H., and Santo, G. S. 1995. Introgression of resistance to
Columbia and northern root-knot nematodes from Solanum bulbocastanum into cultivated potato. Euphytica 83:71–78.
Brown, C. R., Mojtahedi, H., and Bamberg, J. 2004. Evaluation of Solanum fendleri as a source of resistance to Meloidogyne chitwoodi. American Journal of Potato
Research 81:415–419.
Brown, C. R., Mojtahedi, H., Santo, G. S., and Williamson, V. M. 1997. Effect of the Mi gene in tomato on reproductive factors of Meloidogyne chitwoodi and M. hapla.
Journal of Nematology 29:416–419.
Brown, C. R., Mojtahedi, H., Zhang, L. H., and Riga, E. 2009. Independent resistant reactions expressed in root and tuber of potato breeding lines with introgressed resistance to Meloidogyne chitwoodi. Phytopathology 99:1085–1089.
240
Brown, C. R., Yang, C. P., Kwiatkowski, S., and Adiwilaga, K. D. 1991b. Insert copy number, chromosome-number, pollen stainability, and crossability of
Agrobacterium-transformed diploid potato. American Potato Journal 68:317–330.
Brown, C. R., Zhang, L., and Mojtahedi, H. 2014. Tracking the RMc1 gene for resistance to race 1 of Columbia root-knot nematode (Meloidogyne chitwoodi) in three
Mexican wild potato species with different ploidies. American Journal of Potato
Research 91:180–185.
Byrd, D. W., Kirkpatrick, T., and Barker, K. R. 1983. An improved technique for clearing and staining plant tissue for detection of nematodes. Journal of Nematology
15:142–143.
Davies, L. J., Brown, C. R., and Elling, A. A. 2015. Calcium is involved in the
RMc1(blb)-mediated hypersensitive response against Meloidogyne chitwoodi in potato.
Plant Cell Reports 34:167-177.
Dinh, P. T. Y., Brown, C. R., and Elling, A. A. 2014a. RNA interference of effector gene Mc16D10L confers resistance against Meloidogyne chitwoodi in Arabidopsis and potato. Phytopathology 104:1098–1106.
Dinh, P. T. Y., Zhang, L., Brown, C. R., and Elling, A. A. 2014b. Plant-mediated
RNA interference of effector gene Mc16D10L confers resistance against Meloidogyne chitwoodi in diverse genetic backgrounds of potato and reduces pathogenicity of nematode offspring. Nematology 16:669–682.
Elling, A. A. 2013. Major emerging problems with minor Meloidogyne species.
Phytopathology 103:1092–1102.
241
Elling, A. A. and Jones, J. T. 2014. Functional characterization of nematode effectors in plants. Pp. 113–124 in P. Birch, J. Jones, J. I. B. Bos, eds. Plant-pathogen interactions: Methods and Protocols. Methods in Molecular Biology vol. 1127. Berlin,
Germany: Springer Verlag.
Finley, A. M. 1981. Histopathology of Meloidogyne chitwoodi (Golden et al.) on
Russet Burbank potato. Journal of Nematology 13:486–491.
Fire, A., Xu, S., Montgomery, M. K., Kostas, S. A., Driver, S. E., and Mello, C. C.
1998. Potent and specific genetic interference by double-stranded RNA in
Caenorhabditis elegans. Nature 391:806–811.
Gebhardt, C. and Valkonen, J. P. T. 2001. Organization of genes controlling disease resistance in the potato genome. Annual Review of Phytopathology 39:79–102.
Goggin, F. L., Jia, L., Shah, G., Hebert, S., Williamson, V. M., and Ullman, D. E.
2006. Heterologous expression of the Mi-1.2 gene from tomato confers resistance against nematodes but not aphids in eggplant. Molecular Plant-Microbe Interactions 19:
383–388.
Hammond, S. M., Caudy, A. A., and Hannon, G. J. 2001. Post-transcriptional gene silencing by double-stranded RNA. Nature Reviews Genetics 2:110–119.
Huang, G., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S. 2006a.
Engineering broad root-knot resistance in transgenic plants by RNAi silencing of a conserved and essential root-knot nematode parasitism gene. Proceedings of the
National Academy of Sciences USA. 103:14302–14306.
242
Huang, G. Dong, R., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S.
2006b. A root-knot nematode secretory peptide functions as a ligand for a plant transcription factor. Molecular Plant-Microbe Interactions 19:463–470.
Hussey, R. S. and Barker, K. R. 1973. A comparison of methods of collecting inocula of Meloidogyne spp., including a new technique. Plant Disease Reporter
57:1025–1028.
Ingham, R. E., Hamm, P. B., Williams, R. E., and Swanson, W. H. 2000. Control of Meloidogyne chitwoodi in potato with fumigant and non-fumigant nematicides.
Journal of Nematology (Suppl.) 32:556–565.
Janssen, G. J. W., von Norel, A., Verkeerk-Bakker, B., and Janssen, R. 1996.
Resistance to Meloidogyne chitwoodi, M. fallax, and M. hapla in wild tuber-bearing
Solanum spp. Euphytica 92:287–294.
Kyndt, T., Ji, H., Vanholme, B., and Gheysen, G. 2013. Transcriptional silencing of RNAi constructs against nematode genes in Arabidopsis. Nematology 15:519–528.
Lilley, C. J., Davies, L. J., and Urwin, P. E. 2012. RNA interference in plant parasitic nematodes: a summary of the current status. Parasitology 139:630–640.
Mitchum, M. G., Hussey, R. S., Baum, T. J., Wang, X., Elling, A. A., Wubben, M., and Davis, E. L. 2013. Nematode effector proteins: An emerging paradigm of parasitism. New Phytologist 199:879–894.
Sanchez-Puerta, M. V. and Masuelli, R. W. 2011. Evolution of nematode- resistant Mi-1 gene homologs in three species of Solanum. Molecular Genetics and
Genomics 285:207–218.
243
Sheerman, S. and Bevan M. W. 1988. A rapid transformation method for
Solanum tuberosum using binary Agrobacterium tumefaciens vectors. Plant Cell
Reports 7:13–16.
Sikora, R. A. and Fernández, J. 2005. Nematode parasites of vegetables. Pp.
319–392 in M. Luc, R. A. Sikora, J. Bridge, eds. Plant parasitic nematodes in subtropical and tropical agriculture. Wallingford, UK: CAB International.
Wang, C., Lower, S., and Williamson, V. M. 2009. Application of pluronic gel to the study of root-knot nematode behaviour. Nematology 11:453–464.
Williamson, V. M. and Kumar, A. 2006. Nematode resistance in plants: The battle underground. Trends in Genetics 22:396–403.
Zhang, F., Peng, D., Ye, X., Yu, Z., Hu, Z., Ruan, L., and Sun, M. 2012. In vitro uptake of 140 kDa Bacillus thuringiensis nematicidal crystal proteins by the second stage juvenile of Meloidogyne hapla. PLoS ONE 7(6):e38534.
244
6. 8. TABLE
TABLE 1. Reproductive efficiency of Meloidogyne chitwoodi isolate ‘WAMC1’ on wild type and 16D10i-2 RNAi transgenic ‘Russet Burbank’ potato lines at 55 days after inoculation.
Line R = Pf/Pi ± SEa Line R = Pf/Pi ± SEa
RB 82.4 ± 13.14 D20 43.45* ± 8.16
D1 3.76* ± 0.50 D21 0.22* ± 0.16
D2 7.71* ± 2.13 D24 11.31* ± 3.27
D3 35.66* ± 12.89 D25 4.69* ± 0.93
D5 31.54* ± 13.66 D26 28.24* ± 9.34
D7 24.96* ± 5.93 D27 37.86* ± 9.08
D9 2.15* ± 0.93 D32 6.22* ± 1.47
D11 20.75* ± 6.39 D33 4.38* ± 0.86
D14 34.28* ± 3.34 D34 22.49* ± 17.81
D16 70.08 ± 4.45 D35 64.67 ± 21.65
D17 8.8* ± 3.34 D36 2.84* ± 1.47
D18 46.22* ± 6.00
aNematode reproduction R is shown as the ratio of the number of eggs at final harvest Pf to the initial egg inoculum Pi, (R = Pf/Pi) with SE.
RB = ‘Russet Burbank’; * = significant differences compared to RB wild type according to Student’s t-test (P < 0.05).
245
6. 9. FIGURES
FIG. 1. Southern blot showing copy numbers of the 16D10i-2 RNAi transgene in transformed potato lines and controls. For each plant line, 15 µg DNA was digested with
50 U Xba I and separated on a 0.8% agarose gel. 82-4, PA99N82-4 advanced breeding line; RB, wild type ‘Russet Burbank’; E34, empty vector control; D1 to D36, 16D10i-2 transgenic lines.
246
FIG. 2. Number of Meloidogyne chitwoodi ‘WAMC1’ eggs per plant in wild type and transgenic RNAi potato lines at 55 days after inoculation. RB, wild type ‘Russet
Burbank’. Each bar represents the mean of five plants per independent line with SE.
Asterisk indicates statistically significant differences compared to RB controls using a
Student’s t-test (P < 0.05).
247
FIG. 3. Northern blot for 16D10i-2 RNAi transgene. A. Total RNA loading control.
B. U6 small nuclear RNA (snRNA) loading control. C. 16D10i-2-specific small RNA
(smRNA) using probe 16D10 (Dinh et al., 2014a). 82-4, PA99N82-4 advanced breeding line; RB, wild type ‘Russet Burbank’; E34, empty vector control; D21, 16D10i-2 transgenic line D21.
248
FIG. 4. Reproductive success of Meloidogyne spp. in potato lines with and without
16D10i-2 RNAi transgene. Each bar represents the mean of ten plants per independent line and time point with SE. Letters indicate statistically significant differences using a
Student’s t-test (P < 0.05). 82-4, PA99N82-4 advanced breeding line; RB, wild type
‘Russet Burbank’; E34, empty vector control; D21, 16D10i-2 transgenic line D21. A and
249
B. Reproductive success of nematodes as number of egg masses per plant and per gram root fresh weight at 35 days after inoculation (DAI). C and D. Reproductive success of nematodes as number of eggs per plant and per gram root fresh weight at
55 DAI.
250
FIG. 5. Egg masses of Meloidogyne spp. in roots of potato lines with and without
16D10i-2 RNAi transgene. Potato plants were inoculated with nematode eggs and developing egg masses from mature females were stained 35 days after inoculation with phloxine B. A and B. Egg masses of M. incognita and M. javanica, respectively, in roots of 82-4, PA99N82-4 advanced breeding line. C and D. Egg masses of
M. incognita in roots of RB, wild type ‘Russet Burbank’ and E34, empty vector control plants, respectively. E to H. Egg masses of M. incognita, M. arenaria, M. javanica, and
M. hapla, respectively, in D21, 16D10i-2 transgenic line D21. Scale bar = 1 mm.
251
FIG. 6. Attraction and invasion of Meloidogyne incognita J2 to potato roots with and without 16D10i-2 RNAi transgene. A. Number of infective J2 touching the 10-mm terminal end of potato roots with and without 16D10i-2 RNAi transgene at 6 hr after inoculation. B. Number of parasitic J2 invaded potato roots with and without 16D10i-2
RNAi transgene at 4 days after inoculation (DAI). Each bar represents the mean of 15 root tips per independent line with SE. Student’s t-test did not indicate significant differences (P > 0.05). 82-4, PA99N82-4 advanced breeding line; RB, wild type ‘Russet
Burbank’; E34, empty vector control; D21, 16D10i-2 transgenic line D21. C–F. Attraction of M. incognita infective J2 to representative root tips at 6 hours after inoculation in potato lines 82-4, RB, E34, and D21, respectively. G–J. Invasion of M. incognita parasitic J2 in representative roots at 4 DAI after acid fuchsin staining in potato lines 82-
4, RB, E34, and D21, respectively. Scale bar = 100 µm.
252
CHAPTER SEVEN
CONCLUSIONS
This dissertation focused on interactions between host plants and plant parasitic nematodes. At cellular and subcellular levels, a novel and nondestructive technique was developed to observe the entire life cycle of nematodes in planta. Normally, developing nematodes and associated feeding sites are surrounded by many layers of root cells, making it very difficult to directly observe plant-nematode interactions. Recent advances in video-enhanced contrast light microscopy have allowed tracing the behavior of nematodes in A. thaliana roots in agar under axenic conditions. However, some obstacles such as adverse effects of axenic conditions and nematode surface- sterilization, and the instability of fluorescent dyes throughout the entire life cycle still have not been overcome (4, 11, 12).
In this work, the system of microscopy rhizosphere chambers (ROC) provided more natural conditions than axenic media for observing both the growth of A. thaliana and nematode parasitism. With a stable fluorescent stain, PKH26, this technique enabled the entire process of nematode pathogenesis to be observed without tissue destruction. The feeding sites, nematode oogenesis and morphological features of
P. penetrans, H. schachtii and M. incognita were visualized in ROC for up to 27 days after inoculation. In addition, the changes in peroxisome abundance in response to
M. chitwoodi infection as observed by this technique mirrored the transcriptional changes of peroxisome-specific genes reported previously (2, 9, 14). The ROC system not only overcame the inherent challenges in observing nematode behaviors, but has
253 the potential to facilitate the study of root responses to nematode infection and infection by other pathogens such as bacteria, fungi and oomycetes.
In addition to these in vivo observations, host-nematode interactions were studied at the molecular level through the functional characterization of the highly conserved RKN 16D10 effector. The 16D10 was first cloned in M. incognita and found to be highly conserved in other RKN species: M. hapla, M. javanica, and M. arenaria
(8). In this study, an ortholog of the 16D10, Mc16D10L, was cloned in M. chitwoodi suggesting that 16D10 has an essential role in this plant-nematode interaction. To successfully parasitize host plants, RKN modifies host defenses and the division of vascular root cells in order to establish feeding sites. Although cellulose-binding protein, an M. incognita effector, can elongate host roots (7), in this study, overexpressed
16D10 A. thaliana roots were not significantly longer than control roots. However, overexpression of 16D10 increased the number of metaxylem root cells. Induction of metaxylem division could assist M. incognita to form feeding sites. The role of the
16D10 effector in regulation of host defense was not consistent in this study.
This dissertation also demonstrated the potential of using highly conserved
16D10 as a target of RNA interference (RNAi) for RKN control. Plant-mediated 16D10
RNAi was able to silence the Mc16D10L to significantly reduce M. chitwoodi race 1 reproduction by up to 71% in A. thaliana and potato plants (Solanum tuberosum cvs
Russet Burbank and Désirée). This is the first time that the RNAi construct was stably transferred into a crop, potato, to manage RKN parasitism. Because RNAi strategy does not introduce foreign proteins, such as the expressed nematicidal peptides (cystatins and neurotransmitter antagonists), it does not cause food allergies (1). The RNAi
254 strategy also overcomes many disadvantages of traditional breeding such as time consumption, labor costs, and incompatibility barriers (6).
This report also proves for the first time that the silencing effect of 16D10 RNAi can be transmitted to M. chitwoodi offspring as significantly reduced pathogenicity of the nematode offspring was observed on non-RNAi plants. This result is in accordance with the inheritance of RNAi in other nematode taxa, i.e., Caenorhabditis elegans and
M. javanica, giving further evidence that this genetic mechanism is conserved (3, 5, 13).
The 16D10 RNAi effect is assumed to be systemic and heritable in M. chitwoodi. Since a 16D10 silencing Russet Burbank line, D21, was not only resistant to M. chitwoodi; but also found to be resistant to M. incognita, M. javanica, M. arenaria and M. hapla (up to
90% reproductive reduction). Moreover, the 16D10 silencing effect was combined with an M. chitwoodi resistant gene, RMc1(blb), by introducing the 16D10 RNAi construct into
PA99N82-4 potato (a resistant breeding line). 16D10 RNAi PA99N82-4 lines were not only resistant to M. chitwoodi race 1; but also had approximately 50% decrease in the reproduction of M. chitwoodi pathotype Roza, which breaks RMc1(blb).
This study showed that plant-mediated RNAi can be exploited to develop an efficient tool to control RKN in potato as well as other important crops. The RNAi approach had many advantages, such as providing a broad range resistance to RKN species in a diverse potato germplasm and the inheritance of 16D10 RNAi through the nematode offspring. In combination with other control tools, such as applications of resistant genes, green manures, crop rotations, and microbial antagonists, RNAi strategy can reduce the amount of synthetic nematicides used to successfully manage
RKN and protect crop production.
255
LITERATURE CITED
1. Atkinson, H. J., Lilley, C. J., and Urwin, P. E. 2012. Strategies for transgenic
nematode control in developed and developing world crops. Curr. Opin.
Biotechnol. 23:251-256.
2. Barcala, M., García, A., Cabrera, J., Casson, S., Lindsey, K., Favery, B., García-
Casado, G., Solano, R., Fenoll, C., and Escobar, C. 2010. Early transcriptomic
events in microdissected Arabidopsis nematode-induced giant cells. Plant J.
61:698-712.
3. Gleason, C. A., Liu, Q. L., and Williamson, V. M. 2008. Silencing a candidate
nematode effector gene corresponding to the tomato resistance gene Mi-1 leads
to acquisition of virulence. Mol. Plant-Microbe Interact. 21:576-585.
4. Goto, D. B., Fosu-Nyarko, J., Sakuma, F., Sadler, J., Flottman-Reid, M., Uehara,
T., Kondo, N., Yamaguchi, J., and Jones, M. G. K. 2010. In planta observation of
live fluorescent plant endoparasitic nematodes during early stages of infection.
Nematol. Res. 40:15-19.
5. Grishok, A., Tabara, H., and Mello, C. C. 2000. Genetic requirements for
inheritance of RNAi in C. elegans. Science 287:2494-2497.
6. Hermsen, J. G. T. 1994. Introgression of genes from wild species, including
molecular and cellular approaches. Pages 515-538 in: Potato genetics., J. E.
Bradshaw and G. R. Mackay, eds. CAB International, Wallingford, UK.
7. Hewezi, T., Howe, P., Maier, T. R., Hussey, R. S., Mitchum, M. G., Davis, E. L.,
and Baum, T. J. 2008. Cellulose binding protein from the parasitic nematode
256
Heterodera schachtii interacts with Arabidopsis pectin methylesterase:
cooperative cell wall modification during parasitism. Plant Cell 20:3080-3093.
8. Huang, G. Z., Dong, R. H., Allen, R., Davis, E. L., Baum, T. J., and Hussey, R. S.
2006. A root-knot nematode secretory peptide functions as a ligand for a plant
transcription factor. Mol. Plant-Microbe Interact. 19:463-470.
9. Jammes, F., Lecomte, P., Almeida-Engler, J., Bitton, F., Martin-Magniette, M. L.,
Renou, J. P., Abad, P., and Favery, B. 2005. Genome-wide expression profiling
of the host response to root-knot nematode infection in Arabidopsis. Plant J.
44:447-458.
10. Jaouannet, M., Magliano, M., Arguel, M. J., Gourgues, M., Evangelisti, E., Abad,
P., and Rosso, M. N. 2013. The root-knot nematode calreticulin Mi-CRT is a key
effector in plant defense suppression. Mol. Plant-Microbe Interact. 26:97-105.
11. McElwee, M. K., and Freedman, J. H. 2011. Comparative toxicology of
mercurials in Caenorhabditis elegans. Environ. Toxicol. Chem. 30:2135-2141.
12. Sijmons, P. C., Grundler, F. M. W., Mende, N., Burrows, P. R., and Wyss, U.
1991. Arabidopsis thaliana as a new model host for plant-parasitic nematodes.
Plant J. 1:245-254.
13. Vastenhouw, N. L., Brunschwig, K., Okihara, K. L., Müller, F., Tijsterman, M., and
Plasterk, R. H. A. 2006. Long-term gene silencing by RNAi. Nature 442:882-882.
14. Vercauteren, I., Van Der Schueren, E., Van Montagu, M., and Gheysen, G. 2001.
Arabidopsis thaliana genes expressed in the early compatible interaction with
root-knot nematodes. Mol. Plant-Microbe Interact. 14:288-299.
257
APPENDIX
The involvement of the Meloidogyne incognita effector 16D10 in suppression of hypersensitive response (HR) in Nicotiana benthamiana
Methods
To test whether 16D10 suppressed the HR of the plant defense system, 16D10 mature peptide was transiently expressed with HR triggers in N. benthamiana. HR triggers are resistant and avirulent proteins that are involved in the pathogen recognition and the defense response of host plants. Six HR triggers, including Pto(Y207D), RPS2,
AvrRpt2, a combination of ATR13Δ41aa and RPP13, a combination of Gpa2 and RBP-
1, and a combination of CP and Rx2, were used to initiate HR in N. benthamiana. The
16D10 mature peptide was cloned in pEARLYGATE201 under control of the 35S promoter. Empty pEARLYGATE201 vector was used as a control. Agrobacterium tumefaciens (GV3101) carrying pEARLYGATE-16D10 or empty vector control was infiltrated into leaves one day before or co-infiltrated on the same day with
A. tumefaciens carrying the selected HR triggers.
Pto(Y207D) is a mutant of Pto kinase R (resistant) gene cloned from tomato against Pseudomonas syringae pv tomato (Pst) (Rathjen et al. 1999). RPS2, an R protein of Arabidopsis thaliana, interacts with RIN4 to recognize avirulent protein
AvrRpt2 from Pst and causes HR in A. thaliana (Day et al. 2005). HR also is triggered in
N. benthamiana by RPS2 or AvrRpt2 (Day et al. 2005). Moreover, the HR response in
N. benthamiana also can be triggered by co-infiltrating both RPP13, an R gene of
A. thaliana, and ATR13Δ41aa, an avirulence gene of Hyaloperonospora parasitica
(Rentel et al. 2008). The potato R protein, Gpa2 recognizes RBP-1 protein of Globodera
258 pallida to trigger HR in N. benthamiana (Sacco et al. 2009). Similar to Gpa2, potato Rx2 recognizes the coat protein CP of Potato virus X to cause HR (Bendahmane et al.
2000).
Transient expression of 16D10 and HR triggers in N. benthamiana by infiltrating the A. tumefaciens strains followed the protocol described by Sessa and colleagues
(2000). A. tumefaciens strains were cultured at 30ºC for 24 h in Luria-Bertani broth with appropriate antibiotics. Each A. tumefaciens strain was then transferred to induction medium [50 mM MES pH 5.6, 1.7 mM NaH2PO4, 20 mM NH4Cl, 2 mM KCl, 1.2 mM
MgSO4, 17 µM FeSO4, 70 µM CaCl2, 0.5% (w/v) glucose and 200 µM acetosyringone] for 16 h, and then diluted in infiltration solution (10 mM MES and 200 µM acetosyringone) to achieve a final concentration of 0.5 at OD600. Equal ratios of the A. tumefaciens strains were mixed before co-infiltration. For each combination of pEARLYGATE-16D10 and HR triggers, 14 N. benthamiana leaves were infiltrated/co- infiltrated. Four days after infiltration/co-infiltration, HR were photographed with a
Fujifilm camera (Finepix SL260).
Results
When pEARLYGATE-16D10 was infiltrated one day before the HR triggers, Pto, a combination of GPa2 and RBP-1, and a combination of CP and Rx2 still caused HR in all of the infiltrated N. benthamiana leaves. In contrast, the HR caused by RPS2,
AvrRpt2 and a combination of ATR13Δ41aa and RPP13 was prevented in about 50% of infiltrated N. benthamiana leaves that were infiltrated with pEARLYGATE-16D10 one day before these triggers (representative leaf shown in Fig. 1A). The other 50% of the leaves (about 7) still showed HR even though 16D10 was expressed one day prior to
259 exposing to the trigger (representative leaf shown in Fig. 1B). Importantly, 16D10 did not suppress HR in N. benthamiana leaves when pEARLYGATE-16D10 was co- infiltrated with all of the selected HR triggers.
.
A B
3 3 2 2
2 3 1 2
1 1 3 1
Figure 1: The effect of 16D10 in the hypersensitive responses (HR) of
Nicotiana benthamiana
Agrobacterium tumefaciens carrying pEARLYGATE-16D10 was infiltrated into a total of 14 N. benthamiana leaves one day before A. tumefaciens carrying the RPS2 trigger. The HR caused by RPS2 was suppressed by 16D10 (as shown in Fig. 1A) in 7 infiltrated leaves while in the other 7 leaves, 16D10 did not prevent HR (as shown in
Fig. 1B). 1: Infiltration of RPS2 only; 2: Infiltration of empty pEARLYGATE then RPS2 one day later; 3: Infiltration of pEARLYGATE-16D10 then RPS2 one day later.
260
Literature cited
Bendahmane, A., Querci, M., Kanyuka, K., and Baulcombe, D. C. 2000. Agrobacterium
transient expression system as a tool for the isolation of disease resistance
genes: application to the Rx2 locus in potato. Plant J. 21:73-81.
Day, B., Dahlbeck, D., Huang, J., Chisholm, S. T., Li, D., and Staskawicz, B. J. 2005.
Molecular basis for the RIN4 negative regulation of RPS2 disease resistance.
Plant Cell 17:1292-1305.
Rathjen, J. P., Chang, J. H., Staskawicz, B. J., and Michelmore, R. W. 1999.
Constitutively active Pto induces a Prf-dependent hypersensitive response in the
absence of avrPto. EMBO J. 18:3232-3240.
Rentel, M. C., Leonelli, L., Dahlbeck, D., Zhao, B., and Staskawicz, B. J. 2008.
Recognition of the Hyaloperonospora parasitica effector ATR13 triggers
resistance against oomycete, bacterial, and viral pathogens. Proc. Natl. Acad.
Sci. U.S.A 105:1091-1096.
Sacco, M. A., Koropacka, K., Grenier, E., Jaubert, M. J., Blanchard, A., Goverse, A.,
Smant, G., and Moffett, P. 2009. The cyst nematode SPRYSEC protein RBP-1
elicits Gpa2-and RanGAP2-dependent plant cell death. PLoS Pathog.
5:e1000564.
Sessa, G., D'Ascenzo, M., and Martin, G. B. 2000. Thr38 and Ser198 are Pto
autophosphorylation sites required for the AvrPto-Pto-mediated hypersensitive
response. EMBO J. 19:2257-2269.
261